Medical apparatus for breath detection

ABSTRACT

A medical apparatus ( 10 ) for detecting a predetermined component of breath and producing a breath-component signal over a measurement time. Such breath-component signal may be related to a fat metabolism indicator.

RELATED APPLICATIONS

This application is a continuation of U.S. application Ser. No.10/492,953, which is the National Stage of International Application No.PCT/US02/36027, filed Nov. 8, 2002 (title: HAND-HELD MEDICAL APPARATUS),which claims the benefit under 35 U.S.C. 119(e) of U.S. ProvisionalApplication No. 60/332,349, filed on Nov. 9, 2001, each of the foregoingapplications being hereby fully incorporated by reference herein.

FIELD OF THE INVENTION

This invention relates generally to a medical apparatus for breathdetection. More particularly, the invention relates to a medicalapparatus for analyzing acetone in exhaled breath.

BACKGROUND

Diabetes is a chronic disease affecting many organs and body functions.The disease is caused either by a lack of the hormone insulin or by thebody's inability to use insulin. Diabetes is the most common endocrinedisorder. In the United States, for instance, as many as fifteen millionpersons have been diagnosed with diabetes mellitus, and it has beenestimated that an additional ten million may have the disease withoutdiagnosis. Although there is no cure, most cases can now be controlledadequately by a combination of medication and life style modification,including exercise, diet and weight loss.

Unfortunately, many people with diabetes have difficulty coping with theconstraints that the disease puts on their lives. People find itdifficult to lose weight, to maintain weight loss, to exerciseregularly, to regularly take drugs, or to self-administer tests forblood glucose levels. In general, users do not receive sufficientpositive support for their efforts and can become discouraged. Theyexperience “diabetes burn-out”, a feeling of hopelessness orpowerlessness that contributes to abandoning efforts to manage theirdisease. See, for example, Diabetes Burnout, What to Do When You Can'tTake It Anymore, W. H. Polonsky, 1999, American Diabetes Association.

People who are simply overweight or obese can experience similarbarriers as those experienced by individuals managing diabetes, whenattempting to control their diet and weight. Weight loss is bothdifficult to achieve and to sustain. Preferably, for weight loss,caloric intake should be reduced to produce an energy deficit of about300-1000 Calories daily, which usually results in the loss of about onehalf to two pounds of body weight per week (NIH Guidelines, 1998).

The relatively slow rate prescribed for traditional weight loss makesthe measurement of progress to goals difficult to track. Coupled withthe slow rate of weight loss, factors such as daily variation in watercontent of the body, poor sensitivity of most scales, and slight weightgain attributable to contemporaneous improvement in muscle tone fromexercise, can mask the progress being made. Many people, by contrast,expect rapid, dramatic changes in their condition. Still others expectfailure and find this belief confirmed by the slow rate of change intheir health. An accurate, rapid feedback mechanism is needed to helpusers sustain changes in their life style which will lead to sustainedweight loss.

The potential for the use of exhaled breath as a diagnostic tool haslongbeen recognized. Hippocrates taught the physician to be aware of thesmell of the user's breath, as a clue to the user's condition. In 1784Antoine Lavoisier and Pierre Laplace analyzed breath of a guinea pig,finding that an animal inhales oxygen and exhales carbon dioxide. Thiswas the first direct evidence that the body uses a combustion process toobtain energy from food. Since that time, as many as 200 compounds havebeen detected in human breath, some of which have been correlated withvarious diseases.

It is known that a person exhales acetone in the breath when the body isin a condition of energy deficit, that is, when the body is using moreenergy than it is taking in through food or beverages. Ketosis is,therefore, an immediate measurable indication that a person issuccessfully maintaining a reducing diet. See, for example, Samar K.Kundu et al., “Breath Acetone Analyzer: Diagnostic Tool to MonitorDietary Fat Loss”, Clin. Chem., Vol. 39, No. 1, pp. 87-92 (1993).

Detection apparatus for breath components employ varying technologies.Infrared light has been used to measure breath alcohol content by BowidsU.S. Pat. No. 5,422,485 and Paz U.S. Pat. No. 5,515,859. Sauke et al.U.S. Pat. No. 5,543,621 used a laser diode spectrometer. Other types oflasers and absorption spectroscopes have been used includingcavity-ringdown spectroscopy. See, for example “AbsorptionSpectroscopes: From Early Beginnings to Cavity-Ringdown Spectroscopy” B.A. Paldus and R. N. Zare, American Chemical Society Symp. Ser. (1999),Number 720, pp. 49-70. Other techniques include gas-liquidchromatography (“GC”), mass spectrometry, coupled GC-Mass Spectroscopy,electrochemistry, colorimetry, chemi-luminescence, gas biosensors, andchemical methods. See, for example, “The Diagnostic Potential of BreathAnalysis”, Antony Manolis, Clinical Chemistry, 29/1 (1983) pp. 5-15, and“Technology Development in Breath Microanalysis for Clinical Diagnosis”,Wu-Hsum Cheng, et al., J. of Laboratory and Clinical Medicine, 133 (3)218-228 March, 1999. Among the chemical sensors are so-called electronicnoses, which rely on an array of detectors to recognize patterns ofphysical or chemical characteristics to identify components. Thesesensors may rely, for example, on conductive polymers, surfaceacoustical wave generators, metal oxide semiconductors, fluorescence orelectrochemical detection. Such sensors are commercially available fromCyrano Sciences, Pasadena, Calif., for example, and their use indetecting medical conditions such as pneumonia, halitosis and malignantmelanoma has been suggested. Many of these technologies are complex,expensive and difficult to calibrate, and have not been economicallyadapted for individual health care use, let alone portable, hand-heldanalysis.

Medical apparatus for individual health care use have been disclosed. Ithas been suggested that self-administered breath alcohol tests could beused (See, Brown et al. U.S. Pat. No. 5,303,575) by multiple individualsat bars or other locations where alcoholic beverages are served todetect a predetermined level of breath alcohol.

WO 01/63277 and U.S. Patent Application Publication 2002-0007249-A1,herein fully incorporated by reference, disclose a personal computerbreath analyzer for health-related behavior modification. In thedisclosed systems, the user introduces his or her breath into ananalyzer. A computer connected to the analyzer receives abreath-component signal from the sensor and converts the signal to asecond signal. The disclosed systems disclose detection of acetone inbreath for the detection of weight loss. However, the disclosed systemsare not optimized for portable use.

U.S. patent application 2001-0031913-A1 discloses a home health careservice for the monitoring of home health care users. This publicationdiscloses the measurement of an analyte in urine, using a device thatmay be interposed in a toilet. The device detects the presence of achemical component in the urine of the user, and generates an electricalsignal that is transmitted to an Internet-based health care center. Thedisclosed system has not been optimized for portable use. The use of abreath-analyzing biosensor is not disclosed.

U.S. patent application 2001-0056328-A1 discloses a system forcommunications between a biosensor apparatus and a personal dataassistant. The use of a breath-analyzing biosensor is not disclosed. Inaddition, the system requires the user to have a personal dataassistant, which may be viewed by the user as having more functionalitythan is required, and thus, as a source of unwanted expense.

Heath care practitioners and users would find advantage in anon-invasive, and-held, cost-effective system for the real-timemonitoring of fat metabolism.

BRIEF DESCRIPTION OF THE FIGURES

FIG. 1 is a schematic diagram of a hand-held medical apparatusincorporating an electrochemical biosensor for the detection of acetonein breath.

FIG. 2 is a schematic diagram of a hand-held medical apparatusincorporating a sensor for the colorimetric detection of acetone inbreath.

FIG. 3 is a flowchart illustrating one means for the operation of thehand-held medical apparatus of FIG. 1.

FIG. 4 is a flowchart providing further detail regarding the operationof the hand-held medical apparatus of FIG. 1.

FIG. 5 is a drawing of the reaction pathways of an embodiment using apreferred enzyme that selectively targets acetone as the pre-determinedbreath component.

DETAILED DESCRIPTION

Unless otherwise expressly noted in this specification, the singularforms “a”, “an” and “the” include plural referents unless the contentclearly dictates otherwise. Thus, for example, reference to “apre-determined breath component” includes mixtures of pre-determinedbreath components. Unless otherwise expressly noted in thisspecification, all technical and scientific terms used herein have themeaning as commonly understood by one of ordinary skill in the art towhich the invention pertains. Unless otherwise expressly defined in thisspecification, the following terms will be used in accordance with theaccompanying definitions.

“Hand-held” means of a size sufficiently small to permit an adult humanuser of the medical apparatus to generally support the medical apparatuson the palm and optionally the adjacent fingers of one of his or herhands. Typical dimensions are less than about three inches (7.62 cm) byfive inches (12.70 cm) by one inch (2.54 cm). Examples of otherhand-held objects include Palm Pilot™, personal data assistants andpocket calculators.

“Inlet” means an opening to the medical apparatus which is incommunication with the sensor, through which a sample of user breath isintroduced into the medical apparatus for detection or measurement of apre-determined breath component by the sensor.

“User breath” means a sample of breath exhaled from the user into abreath inlet of the medical apparatus, which user breath may contain apre-determined breath component.

“Sensor” means a device comprising means for recognizing thepre-determined breath component and means for transducing a signal basedupon such recognition.

“Pre-determined breath component” means a potential component of userbreath to be detected by the sensor, which potential component is aby-product of fat metabolism. Detection or measurement of at least onesuch pre-determined breath component will permit the determination ofwhether or not the user is metabolizing fat at the time that a givensample of user breath is detected by the sensor. One exemplarypre-determined breath component is acetone.

“Breath component signal” means an electrical or optical signalgenerated by the sensor when the sensor is exposed the pre-determinedbreath component.

“Measurement time” means the period of time over which the sensorproduces the breath component signal.

“Sensing electrical circuit” means an electrical circuit adapted forcommunicating the breath component signal from the sensor to the analogto digital converter. The sensing electrical circuit optionally andpreferably includes an amplifier for amplifying the breath componentsignal prior to communication to the analog to digital converter.

“Operative connection” means a connection between two units, objects ordevices which is suitable to permit physical communication or electricalcommunication from one to the other or to permit the two units, objectsor devices to work in concert with one another to achieve apre-determined cooperative effect.

“Analog to digital converter” means a device for converting an analogbreath component signal received from the sensing electrical circuit toa digital signal receivable by the microprocessor.

“Microprocessor” means a computer processor contained on an integratedcircuit chip, preferably including memory and associated circuits.

“Data signal” means a signal derived from the digital signal, which ismeaningful to the user. By way of non-limiting example, data signalsinclude a binary signals (indicating the presence or absence ofdetectable levels of the pre-determined breath component in the userbreath), as well as quantitative signals (indicating the concentrationof the pre-determined breath component in said user breath).

“User fat metabolism indicator” means a numeric, audio, visual,audiovisual and/or tactile signal for display to the user. By way ofexample, the user fat metabolism indicator may be the data signalitself. In the alternative, the user fat metabolism indicator may be anexpression of the data signal, for instance, a display of “detected” or“non-detected” or a symbol therefor, such as a green light (detected) ora red light (not-detected), or a bell (detected) or buzzer(not-detected). Alternatively, the user fat metabolism indicator maysignal whether or not the concentration of the predetermined breathcomponent falls within a pre-designated range (in which case, the userfat metabolism indicator may be, for instance, “recommended level forsafe weight loss”, “lower level than recommended level for effectiveweight loss”, or “higher level than recommended for safe weight loss”).Likewise, the user fat metabolism indicator may indicate progress in aweight control program.

“Electrochemical biosensor” means a sensor which detects electricitygenerated from a biochemical transformation of the pre-determined breathcomponent (or a reaction product or by-product thereof. In a preferredembodiment, said electrochemical biosensor will comprise a disposableelectrode system, which in turn comprises a working electrode, referenceelectrode, and a counter electrode; a physical support; and an enzymethat selectively targets the pre-determined breath component. In oneembodiment, the electrodes will be screen-printed onto the physicalsupport. In one embodiment, the enzyme will be immobilized in a gel orpolymeric medium, which is retained in communication with the electrodeson the surface of the electrochemical biosensor.

“Personal data assistant” means a hand-held computing device havingsuitable power sources and electronics, for example, memory means,software means and display means, which is suitable to receive a datasignal from a microprocessor within the hand-held medical apparatus,store the data signal as stored data, convert the data signal to a userfat metabolism indicator, and display the user fat metabolism indicator(or related information) for observation by the user. The personal dataassistant may be a commercially available personal data assistant, suchas a Palm™, personal data assistant, having a port suitable forproviding a removable operative connection between the personal dataassistant and hand-held medical apparatus. In the alternative, thepersonal data assistant may be a specially adapted device having ahousing, wherein the sensor, suitable power sources and electronics, forexample, memory means, software means and display means, are retainedwithin or are fixedly attached to and/or retained within the housing.

In one embodiment, there is provided a hand-held medical apparatuscomprising:

-   -   a. a housing;    -   b. an inlet for receiving a sample of user breath;    -   c. a sensor for detecting a pre-determined breath component of        said user breath and producing a breath-component signal over a        measurement time;    -   d. a sensing electrical circuit in electrical communication with        said sensor for sensing said breath-component signal, wherein        the magnitude of said breath-component signal is a function of        the concentration of said pre-determined breath component in        said breath sample to be received into said inlet;    -   e. an analog to digital converter in electrical communication        with said sensing electrical circuit for converting said        breath-component signal to a digital signal;    -   f. a microprocessor for processing said digital signal into at        least one of a data signal and a user fat metabolism indicator;        and    -   g. a display in electrical communication with said        microprocessor for displaying said user fat metabolism        indicator.

This embodiment is expected to assist a user in modifying health relatedbehaviors, particularly weight loss. In particular, the inventiveapparatus is expected to provide a measurement that is more reflectiveof the user's recent choices than other typical measurement devices,such as scales, tape measures, and fit of clothing. The inventiveapparatus is expected to enable the user to readily recognize progressmade and the lack thereof, and associate the same with the dietary andexercise choices that he or she has recently made. The portable andhand-held nature of the inventive apparatus is expected to beadvantageous, in that it will accord the user with convenience andprivacy during use.

Various embodiments are now described in connection with theaccompanying figures, wherein like numerals are used to designate likeparts in each drawing. FIG. 1 provides a block diagram of one embodimentof a hand-held medical apparatus. Hand-held medical apparatus 10comprises housing 12. Apparatus 10 has an inlet 14 in communication withconduit 15. In one preferred embodiment, inlet 14 is disposable and/ordetachable from housing 12.

An electrochemical biosensor 16, preferably comprising a workingelectrode, counter electrode and reference electrode, is in electricalcommunication with sensing electrical circuit 17. Sensing electricalcircuit 17 is in electrical communication with an analog to digitalconverter 18. A constant voltage circuit 19 is in electricalcommunication with the sensing electrical circuit 17 and the analog todigital converter 18. A battery, not shown, is used to power thehand-held medical apparatus 10 and, of course, other power sources canbe used such as a converter. The digital signal from the analog todigital converter 18 is communicated to a microprocessor 20. Themicroprocessor 20 is in electrical communication with a liquid crystaldisplay 21 and a personal data assistant 22.

A preferred enzyme system used in the electrochemical biosensor 16 isdescribed in greater detail below. A more general description of theenzyme system is disclosed in U.S. Provisional Patent Application Ser.No. 60/332,349 filed Nov. 9, 2001, herein fully incorporated byreference. Details of suitable electrode designs and electroniccomponents that can be used herein are described in U.S. Pat. Nos.5,571,395 and 5,656,142, herein fully incorporated by reference.

Hand-held medical apparatus 10 can include a sampling device, not shown.The sampling device captures a portion of the user's exhaled breath,preferably alveolar breath from deep within the lungs. The breath samplemay be captured in a chamber or in a trap or both. Generally, traps fallinto three categories: chemical; cryogenic, cold trapping or condensing;and adsorptive. Highly preferred is a trap that utilizes a compressible,porous material, such as open cell polyurethane foam, which can holdwater or buffer solution. The porous material allows the water or buffersolution to be held in a dispersed state with a high surface area. Asthe user blows through the porous material, acetone is partitioned intothe water or buffer solution according to Henry's Law. The porousmaterial can then be compressed to release the water or buffer solutionthat now contains the acetone onto the electrochemical biosensor 16.Alternatively, the water or buffer solution can be conducted out of theporous material by a capillary channel at the tip of the electrochemicalbiosensor 16. A mass air sensor system, not shown, can be positioned inthe conduit 15 to better assure that a sufficient volume of user breathhas been introduced into the inlet 14 of the hand-held medical apparatus10.

Hand-held medical apparatus 10 can also include data storage means, notshown, in electrical communication with the microprocessor 20.Microprocessor 20, for example, can convert a data signal to a user fatmetabolism indicator. Liquid crystal display 21 displays to the user oneor more elements of data, including but not limited to the data signaland the user fat metabolism indicator.

Hand-held medical apparatus 10 can further comprise user input means,not shown, such as a keyboard, mouse, voice recognition device, or anelectronic stylus, through which the user can introduce additionalinformation to microprocessor 20. Apparatus 10 preferably furthercomprises communication means 23, by which information, including butnot limited to the data signal or user fat metabolism indicator, istransmittable to, for example, the personal data assistant 22 and/or toa computer (not shown) for data storage and/or further processing.Communication means 23 include but are not limited to means forestablishing a wired connection, wireless connection, telephonicconnection or Internet connection between microprocessor 20 and anexternal computer. Preferably, such external communication means (aswell as the personal data assistant means themselves) will include dataencryption means for preserving the privacy of individual user data. Aclock, not shown, will preferably be provided and connected to orincorporated in microprocessor 20 or the personal data assistant 22.Personal data assistant 22 can alternatively be located within housing12.

In one preferred embodiment, hand-held medical apparatus 10 willcomprise one or more additional sensors, not shown. These sensors mayinclude an environmental thermometer, a barometer, a hygrometer, orother sensors for determining the condition in which the sample isgiven. The sensors may also include additional user sensors, such as auser thermometer, heart rate or blood pressure sensors. Another sensormight be a camera or voice recognition device to confirm the user'sidentity as well as to record more information on the user's health. Theoutput from such sensors can be communicated to microprocessor 20 and/orthe personal data assistant 22 and optionally stored in data memory. Thepersonal data assistant 22 can optionally be in one or two waycommunication directly or indirectly with a computer, such as by way ofthe Internet.

Hand-held medical apparatus shown in FIG. 1 can be used by first wettingthe electrochemical biosensor 16 and inserting it through the side ofthe housing 12 into the conduit 15, breathing into the inlet 14 for alength of time sufficient to equilibrate the electrode biosensor 16 withacetone from the user breath and then pressing an initiating button. Thesensing electrical circuit applies a voltage to the electrochemicalbiosensor 16 (for example, about 350 millivolts between the working andreference electrodes; determining the electrical current between theworking electrode and the counter electrode) and reads the electricalcurrent between the working electrode and the counter electrode after apredetermined time, for example, after thirty seconds (or morepreferably such current is measured over time as described below, toobtain an integrated signal) as the breath-component signal. The analogto digital converter 18 converts the breath-component signal to adigital signal sent to the microprocessor 20. The microprocessor 20sends a data signal or user fat metabolism indicator to the liquidcrystal display 21 and to the personal data assistant 22. Prior to use,the electrochemical biosensor 16, will preferably be calibrated byexposing it to known concentrations of solutions of the pre-determinedbreath component (for instance, acetone) in water, and deriving astandard curve.

The hand-held medical apparatus will preferably recognize and rejectintroduction of electrochemical biosensors that have already beenexposed to user breath.

FIG. 2 depicts an alternate apparatus embodiment 30. Hand-held medicalapparatus 30 comprises housing 32. Apparatus 30 has adisposable/detachable inlet 34 in communication with transparent conduit35. A disposable enzyme colorimetric biosensor 36 is inserted into thehousing 32 and the conduit 35 as shown. A light source 37 is used todirect light onto the biosensor 36. A light detector 38 is used todetect or diffuse light from the biosensor 36. The light detector 38 isin electrical communication with an amplifier 39. The amplifier 39 is inelectrical communication with a track and hold circuit 40. The track andhold circuit 40 is in electrical communication with an analog to digitalconverter 41. A battery, not shown, is used to power the apparatus 30and, of course, other power sources can be used such as a converter. Theanalog to digital converter 41 is in electrical communication with amicroprocessor 42. The microprocessor 42 is in electrical communicationwith a liquid crystal display 43 and a first computer 44. The firstcomputer 44 is in communication with a second computer 45 by way of anInternet connection.

A preferred enzyme system used in the biosensor 36 is described ingreater detail below and used, for example, with Trinder dye systemcoupled to horseradish peroxidase. A more general description of thecolorimetric enzyme system is disclosed in above referenced U.S.Provisional Patent Application Ser. No. 60/332,349 filed Nov. 9, 2001.Details of suitable electronic circuits and components that can be usedherein are described in U.S. Pat. Nos. 4,935,346 and 5,426,032, hereinfully incorporated by reference. A mass air sensor system, not shown,can be positioned in the conduit 35 to better assure that a propervolume of user breath has been introduced into the inlet 34 of theapparatus 30. Alternatively, the enzyme system used in the biosensor 36can be fluorescence based (in which case the light source 37 is theexcitation light for the fluorescence) or the enzyme system used in thebiosensor 36 can be chemiluminescence based (in which case the lightsource 37 is not used).

FIG. 3 is a flowchart illustrating one means for the operation ofmedical apparatus 10 of FIG. 1. As shown in FIG. 3, at box 100, the userpowers on medical apparatus 10. At box 102, the user selects the desiredaction from a menu, using user input means. Exemplary desired actionsinclude the actions of boxes 200 (making a new measurement of the userfat metabolism indicator), 300 (making an additional measurement usingbiosensor 16), 400 (inputting additional data), 500 (downloading data toor from an external computer), 600 (running a report), or 700 (poweringmedical apparatus 10 off). Upon completion of the selected desiredaction, the user returns to box 102, for selection of an additionaldesired action, repeating the process until the session is completed atbox 700 by the powering off of medical apparatus 10.

FIG. 4 is a flowchart providing further detail regarding box 200 of FIG.3 (take a new measurement of the user fat metabolism indicator).

At box 210, the medical apparatus 10 initializes. At box 220, the useris prompted to breathe into the medical apparatus at inlet 14. Box 220may include means for ascertaining whether the breath sample satisfiespre-determined measurement criteria, and communicating to the user ifsuch pre-determined measurement criteria are not met. Examples ofpre-determined measurement criteria include the volume of user breathintroduced and the duration of time between the preceding measurementand the present measurement (for instance, if it is desired that theuser refrain from making measurements more frequently than atpre-determined intervals).

At box 230, the medical apparatus 10 will determine the concentration ofthe pre-determined breath component, for instance, acetone, in thesample of user breath. Loop 235 indicates a preferred embodiment, inwhich additional samples of user breath are introduced (for apredetermined number of times or until the reading stabilizes. In thisembodiment, the independent measurements of the predetermined breathcomponent may be averaged (or the stabilized measurement determined) andcommunicated to box 240 for calculation of the user fat metabolismindicator.

At box 240, the user fat metabolism indicator is calculated. Thecalculation may involve determining whether or not the concentration ofthe predetermined breath component falls within a pre-designated range(in which case, the user fat metabolism indicator may be, for instance,“recommended level for safe weight loss”, “lower level than recommendedlevel for effective weight loss”, or “higher level than recommended forsafe weight loss”). In this case, the user fat metabolism indicator isdetermined by microprocessor 20.

At box 250, the user fat metabolism indicator will preferably be storedin memory.

At box 260, the user fat metabolism indicator will be displayed to theuser.

Further detail regarding means by which additional user data is inputinto medical apparatus 10 or 30 will now be discussed. In addition tomeasuring a physiologic parameter correlated to a behavior or conditionto be changed (for example, breath acetone as a marker for weight loss)and correlating stored patterns of that parameter, information on thepsychological or emotional state of the user can be obtained. Thisinformation may be either directly obtained from the user or may beinferred from a medical history stored in a computer or both. To acquireinformation directly, the computer may pose a series of questions to theuser. The user may be asked to indicate their perceived state on ascale, for instance. Preferably, the questions are changed from time totime, so that merely routine answers are less likely. Information on theuser's emotional or psychological state may also be inferred from thehistory maintained by the computer. For instance, early enthusiasm for aweight-loss program may be correlated with regular use of the breathanalyzer to detect acetone, and a consistent pattern of acetone levels.Discouragement may be indicated by sporadic and increasingly infrequentuse of the device, coupled with consistently low levels or fluctuatinglevels of detected acetone.

The physiologic parameter and the information on the psychological oremotional state of the user are then correlated to select an appropriateresponse or feedback for the user. For example, adequate levels ofacetone in the breath combined with a feeling of general satisfactionmay produce a response merely acknowledging that the user is in factmeeting his or her goals. Indications of discouragement coupled withadequate physiologic parameter may require more emphatic positivereinforcement to help the user recognize that he or she is makingprogress. A depressed emotional state and poor physiologic measurementsmay require outside intervention. Intervention may include automaticallyalerting a health care provider or a support person or support group sothat personal contact may be made. A connection may be automaticallyinitiated through a communications network as discussed above, forexample, telephone or the Internet system, to the health care provideror support person, reporting the probable need for intervention.

The type of feedback provided to the user may also depend on the user'shistory as recorded by the computer. A process of changing ahealth-related activity or behavior may be viewed as a project or newjob and is characterized by an emotional state which is related to theduration of the project, called herein “an intermediate-term emotionalstate”. Persons undertaking a project generally are observed to be inone of four states or conditions at different times during the project,each state needing a particular type of feed back. A successful projectprogresses through the four phases. A particular user may take more orless time in a particular phase and may, at times, regress to an earlierstate. The four phases may be characterized as a beginning ororientation phase, a dissatisfaction phase, a production or performancephase, and a completion phase. As the project of changing behaviorbegins, the user is usually enthusiastic, but has little realinformation relevant to the change in behavior. For example, the user isexcited about the prospect of improving health by weight loss, butdoesn't know how to prepare appropriate meals in appropriate amounts. Ingeneral, specific, detailed direction is needed in this phase and thecomputer would provide detailed help. Health benefits are not yetapparent to the user.

In the second phase, the health benefits have still not become obvious,and the user may feel discouraged or dissatisfied. This phase needsfeedback that is still detailed but which also includes positivere-enforcement to boost morale. In the case of weight loss, thedetection of acetone components in the breath can provide immediatepositive re-enforcement necessary to help the user through this phase.

In the third phase, physical changes begin to become apparent to theuser. The behavior can be seen to be having the desired effect. Theuser's morale improves and feedback from the system should become lessdetailed and more supportive. In other words, the user's range ofchoices increases as the user becomes accustomed to the changed patternof behavior. Positive re-enforcement is still needed.

In the final phase, the acquired pattern of behavior can be maintainedindefinitely. The user's morale and performance are both high. Detailedinstructions are not needed and would not contribute to maintaining thedesired behavior. Recognition and reward are needed to confirm thesuccessful completion of the changed state. The user maintains the newhabits. In the case of weight loss, for example, acetone is asignificant breath component only during weight loss, when the body isoperating at an energy deficit. When the user is maintaining aparticular weight, measurable levels of acetone may not be detected.

The psychological pattern described above generally extends over theduration of an entire project. In the case of sustained weight loss,this period is usually about a year, comprised of six months of actualweight loss and six months of maintenance to allow the body to acclimateto the lower weight. Dieters and other persons trying to change ahealth-related behavior also experience wide emotional or psychologicalvariation on a short-term basis. The person's need for re-enforcementand support may vary substantially throughout a single day. A recognizedphenomenon in diabetics who are trying to lose weight is the tendency toover eat at the end of the day. Emotional states such as boredom, guilt(for eating “forbidden” foods), and lack of emotional support contributeto this phenomenon. By monitoring the user's emotional state throughoutthe day, additional support or responses can be provided to help theuser cope with the short-term variations that can provide a significantbarrier to successful behavior modification.

In an especially preferred embodiment of the invention, the sensor willinclude an enzymatic system. Suitable enzymes which utilize acetone as asubstrate include secondary alcohol dehydrogenases, acetonemonooxygenases and acetone carboxylases. A more general description ofthese acetone-specific enzyme systems is disclosed in above referencedU.S. Provisional Patent Application Ser. No. 60/332,349 filed Nov. 9,2001.

Referring now to FIG. 5, one preferred secondary alcohol dehydrogenase(S-ADH) for use in an acetone specific enzyme system is NADH-dependentS-ADH isolated from the Gram-negative soil bacterium Xanthobacterautotrophicus strain Py2 (referred to herein as X. autotrophicus Py2 oras X. autotrophicus st. Py2; ATCC deposit number PTA-4779). This enzymecatalyzes the reduction of acetone in the presence of the reducedpyridine nucleotide cofactor NADH to form 2-propanol and NAD⁺.Preferably the S-ADH enzyme reaction is coupled to lactate dehydrogenaseand pyruvate oxidase enzyme activities so that acetone present in theenzyme system is stoichiometrically converted to H₂O₂. H₂O₂ is thenelectrochemically oxidized and detected at the electrode where theanodic current output directly correlates to the concentration ofacetone present. Alternatively, horseradish peroxidase may be added inthe presence electron donors (chromogenic dye reagents) to allowmonitoring of the reaction photometrically.

One preferred acetone monooxygenase for use in an acetone-specificenzyme system is cytochrome P450 acetone monooxygenase, which has beenisolated from mice (Mus musculus). This monooxygenase has been reportedto utilize acetone as a substrate to produce acetol, and is commerciallyavailable from PanVera Corporation (Madison, Wis.). See F. Y. Bondoc, etal., Acetone catabolism by cytochrome P450 2E1: Studies with CYP2E1-nullmice. Biochemical Pharmacology, 58: 461-463 (1999). The enzymeresponsible for this activity in bacteria has not yet been fullycharacterized.

Preferred acetone carboxylases for use in an acetone-specific enzymesystem include acetone carboxylase obtained: from Xanthobacterautotrophicus strain Py2 (see Sluis, M. K. and Ensign, S. A.,Purification and characterization of acetone carboxylase fromXanthobacter strain Py2, PNAS USA, 94: 8456-8462 (1997)); fromRhodobacter capsulatus B10 (see Sluis, M. K. et al., Biochemical,Molecular, and Genetic Analyses of the Acetone Carboxylases fromXanthobacter autotrophicus Strain Py2 and Rhodobacter capsulatus StrainB10, J. Bacteriol., 184(11), 2969-2977 (2002)); and from Rhodococcusrhodochrous B276 (see Clark, D. D. and Ensign, S. A., Evidence for aninducible nucleotide-dependent acetone carboxylase in Rhodococcusrhodochrous B276, J. Bact. 181(9): 2752-2758 (1999)). The Xanthobactergene sequences are available in GenBank (accession number AY055852).,and Rhodobacter capsulatus B10 genes are available on the website of the“Rhodobacter Capsulapedia” sequencing project (Seehttp://rhodol.uchicago.edu). Both the X. autotrophicus Py2 and the R.capsulatus B10 enzymes are (α/β/γ) heterotrimers, sharing approximatelyan 80% overall sequence identity with each other, as well as exhibitingfunctional identity in catalyzing the same reaction with acetone.

A preferred embodiment of the invention provides a breath acetonediagnostic device having one or more acetone-specific enzyme systems. Apreferred use of such a device is in monitoring ketone production in amammal. Acetone-specific enzyme systems are employed in such a way sothat in the presence of acetone, oxidized pyridine nucleotides orhydrogen peroxide are formed as co-products, allowing the reaction bedetected electrochemically. These oxidoreductase enzyme systems include,for example: 1) the secondary alcohol dehydrogenase (S-ADH)-catalyzedreduction of acetone with concomitant NAD(P)H consumption; 2) acetonecarboxylase reaction coupled to β-hydroxybutyrate dehydrogenaseconsumption of NAD(P)H; 3) acetone carboxylase reaction ATP hydrolysiscoupled to NAD(P)H consumption; 4) S-ADH reaction NAD(P)⁺ formationcoupled to H₂O₂ formation; 5) acetone carboxylase reaction ATPhydrolysis coupled to H₂O₂ formation; 6) acetone carboxylase reactioncoupled to β-hydroxybutyrate dehydrogenase NAD(P)⁺ formation coupled toH₂O₂ formation; 7) acetone monooxygenase coupled to NAD(P)H oxidation;and 8) acetone monooxygenase coupled to coupled to H₂O₂ formation. Inall of these enzyme systems, the pyridine nucleotide or hydrogenperoxide is detectable electrochemically. Of course, other detectionmeans known in the art (such as colorimetry, fluorescence,chemiluminescence and calorimetry) can also be utilized.

With regard to some of the terms used herein, NAD(P)⁺ is used herein tomean “either or both of NAD⁺ (nicotinamide adenine dinucleotide,oxidized form) and NADP⁺ (nicotinamide adenine dinucleotide phosphate,oxidized form).” NAD(P)H is used herein to mean “either or both of NADH(nicotinamide adenine dinucleotide, reduced form) and NADPH(nicotinamide adenine dinucleotide phosphate, reduced form).”Nicotinamide adenine dinucleotide is also called3-carbamoyl-1-D-ribofur-anosyl-pyridinium hydroxide 5′-ester withadenosine 5′-pyrophosphate, inner salt. Nicotinamide adeninedinucleotide phosphate is also called3-carbamoyl-1-D-ribofuranosyl-pyridinium hydroxide 5′→5′-ester withadenosine 2′-(dihydrogenphosphate) 5′-(trihydrogen pyrophosphate), innersalt. As used herein, “A_(NNN)” indicates “absorbance measured at NNNnanometers wavelength.” As used herein, “NNN” indicates “extinctioncoefficient measured at NNN nanometers wavelength.”

“Enzyme” as used herein means “catalytically functional biomolecule;”thus any biomolecule that can perform a named catalytic function as itsprimary catalytic activity is considered an enzyme of that name,regardless of other considerations such as origin, native or engineeredstructure, size, etc.

“Platinized carbon,” as used herein, indicates platinum-coated carbon,for example at least partially platinum-coated carbon nanoparticles.

“Photometric,” as used herein, indicates any detection mode in whichphotons are utilized and includes, but is not limited to, colorimetric,spectrometric, spectrophotometric, luminescence-based,chemiluminescence-based, electrogenerated chemiluminescence-based,ioluminescence-based, and fluorescence-based methods.

In order to address certain difficulties associated with subject healthmaintenance, an enzyme-based biosensor has been developed, which enablesthe coupling of enzyme-mediated metabolism of acetone toelectrochemically detectable signals produced via one or more of thesignal mediators. Any acetone-specific enzyme capable of linkage to anelectrochemically detectable co-factor or by-product may be suitable forthe enzyme system of the invention. In a preferred embodiment, anelectrochemical biosensor for detecting acetone in a biological samplecontains at least one acetone-specific enzyme system, and a means fordetecting a product resulting from a reaction between the at least oneacetone-specific enzyme system and acetone in the biological sample. Thedetection means may be either electrochemical or non-electrochemical.

Acetone-Specific Enzymes

A number of enzymes, mainly from bacterial sources, have been describedwhich specifically utilize acetone as a substrate. These enzymes havebeen obtained from and/or characterized in aerobic and anaerobicbacteria that are able to grow using acetone as a sole carbon and energysource.

Acetone may be formed in bacteria by the action of secondary alcoholdehydrogenase (S-ADH), an enzyme that operates in conjunction with oneof two different acetone metabolic pathways: an O₂-dependent (oxygenutilizing) pathway in which the acetone is then oxidized to produceacetol, and a CO₂-dependent (carbon dioxide utilizing) pathway in whichthe acetone is then converted to acetoacetate. The acetone formationreaction catalyzed by S-ADH is freely reversible and normally requires acoenzyme that is typically either NAD(H) or NADP(H). The reduction ofacetone to isopropanol by oxidation of NAD(P)H (the reverse,S-ADH-catalyzed reaction) involves redox chemistry by which acetoneconcentration can be monitored (for example, by means of electrochemicaldetermination of NAD(P)H consumption). A variety of secondary alcoholdehydrogenases have been purified and characterized. Those best studiedare S-ADHs obtained from hydrocarbon oxidizing (that is propaneutilizing) bacteria, which employ O₂-dependent acetone metabolicpathways. S-ADH enzymes have also been isolated from or described inmicroorganisms not associated with hydrocarbon oxidation (that ispropane degradative metabolism). These include methylotrophic bacteriaand yeast, methanogenic Archaea, and fermentative anaerobes. Of theseenzymes, S-ADH from Thermoanaerobium brockii is commercially availableas a heat-treated crude preparation or in purified form (available fromSigma Chemical Co., St. Louis, Mo.). This enzyme is well characterizedand is an NADPH-specific dehydrogenase.

In some propane-oxidizing bacteria, acetone is formed as an intermediatethat is then understood to undergo hydroxylation in an O₂-dependentmono-oxygenase-catalyzed reaction to form acetol (hydroxyacetone).Acetol is then further oxidized to methylglyoxal catalyzed by an acetoldehydrogenase, or is involved in a carbon-carbon cleavage reactionproducing C1 and C2 fragments. Acetone mono-oxygenase, which is apyridine nucleotide-dependent enzyme, provides the necessaryrequirements for electrochemical detection in an acetone biosensor (asdescribed above). Acetone metabolism via acetol as an intermediate hasbeen identified in in vivo studies of acetone-utilizing bacteria. Also,P450 mono-oxygenases have been identified in mammals as using anidentical mechanism (to oxidize acetone to acetol). An acetonemono-oxygenase suitable for use in an acetone-specific enzyme system isa cytochrome P450 acetone mono-oxygenase isolated from mice (Musmusculus). This monooxygenase has been reported as utilizing acetone asa substrate to produce acetol, and is commercially available fromPanVera Corporation (Madison, Wis.). See F. Y. Bondoc et al. Acetonecatabolism by cytochrome P450 2E1: Studies with CYP2E1-null mice.Biochemical Pharmacology, 58: 461-63 (1999). The enzyme responsible forthis activity in bacteria has not yet been fully characterized. Inaddition, acetone mono-oxgenase can be coupled to H₂O₂ generation byincluding a galactose oxidase in the enzyme system; galactose oxidaseoxidizes acetol to form H₂O₂ which can be detected eitherelectrochemically or non-electrochemically.

Mammalian P450 cytochromes containing acetone mono-oxygenase activityand P450 reductase may be prepared from heptatic microsomes. P450acetone mono-oxygenase catalyzes the following hydroxylation reaction:NAD(P)H+H⁺+acetone+O₂→NAD(P)⁺+acetol+H₂O (P450 acetone mono-oxygenase)

P450 monooxygenases are typically comprised of two enzyme componentsincluding a pyridine nucleotide-dependent reductase and an activesite-containing oxygenase component. NAD(P)H provides the necessaryreductant for O₂ activation and incorporation of one oxygen atom intothe aliphatic hydrocarbon substrate. With some P-450 monooxygenases, athird electron transfer component, cytochrome b₅, will stimulateactivity. Acetone-dependent consumption of NAD(P)H by an acetonemono-oxygenase reaction could be monitored electrochemically asdescribed below for secondary alcohol dehydrogenase-coupled and acetonecarboxylase-coupled enzyme systems, as shown in FIG. 1. Alternatively,the reaction could be monitored by following O₂ consumptionelectrochemically, or monitored optically by measuring absorbance orfluorescence of NAD(P)H consumption as described below.

For other bacteria, including both aerobes and anaerobes, acetonemetabolism is has been identified as proceeding by a CO₂-dependentcarboxylation reaction producing acetoacetate. Acetone carboxylase, theenzyme that catalyzes this reaction, has recently been purified tohomogeneity from two bacterial sources. Although acetone carboxylasedoes not catalyze a reaction that is readily detectableelectrochemically, this enzyme has high specificity for acetone and,according to the present invention, can be coupled with other enzymesthat catalyze redox reactions (for example dehydrogenases, oxidases).The feasibility of using coupling enzymes with acetone carboxylase forelectrochemical detection had not been reported prior to thisdisclosure.

Suitable acetone carboxylases for use in an acetone-specific enzymesystem include, but are not limited to, acetone carboxylase obtained:from Xanthobacter autotrophicus strain Py2 (referred to herein as X.autotrophicus Py2 or as X. autotrophicus st. Py2) (see Sluis, M. K. andEnsign, S. A., Purification and characterization of acetone carboxylasefrom Xanthobacter strain Py2, PNAS USA, 94: 8456-8462 (1997)); fromRhodobacter capsulatus B10 (see Sluis, M. K. et al., Biochemical,Molecular, and Genetic Analyses of the Acetone Carboxylases fromXanthobacter autotrophicus Strain Py2 and Rhodobacter capsulatus StrainB10, J. Bacteriol., 184(11):2969-77 (2002)); and from Rhodococcusrhodochrous B276 (see Clark, D. D. and Ensign, S. A., Evidence for aninducible nucleotide-dependent acetone carboxylase in Rhodococcusrhodochrous B276, J. Bact. 181(9):2752-58 (1999)). Xanthobacterautotrophicus strain Py2 was deposited in the American Type CultureCollection (ATCC) on Oct. 29, 2002 under ATCC Accession No. PTA-4779.The ATCC is located at 10801 University Boulevard, Manassas, Va.20110-2209 U.S.A. and may be contacted at P.O. Box 1549, Manassas, Va.20108 U.S.A. This deposit was made in accordance with the requirementsof the Budapest Treaty. The amino acid sequences of the subunits of theX. autotrophicus Py2 acetone carboxylase are set forth in SEQ ID NOs:1,2, and 3; the nucleotide sequences of the genes encoding these subunitsare available in GenBank (See accession number AY055852). The amino acidsequences of the Rhodobacter capsulatus B10 acetone carboxylase gene areset forth in SEQ ID NOs:4, 5, and 6; the nucleotide sequences of thegenes encoding these subunits are available on the website of the“Rhodobacter Capsulapedia” sequencing project (Seehttp://rhodol.uchicago.edu). Both the X. autotrophicus Py2 and the R.capsulatus B10 acetone carboxylase enzymes are alpha/beta/gamma (α/β/γ)heterotrimers, sharing approximately an 80% overall sequence identitywith each other, as well as exhibiting functional identity in catalyzingthe same reaction with acetone.

Acetone-Specific Enzyme Systems

In a preferred embodiment of the invention, a breath acetone diagnosticdevice is provided that contains one or more acetone-specific enzymesystems. A preferred use of such a device is in monitoring ketoneproduction in a mammal. In developing the invention, a number ofoxidoreductase enzyme systems were investigated that, in the presence ofacetone, oxidized pyridine nucleotides as cofactors or produced hydrogenperoxide as a co-product, allowing the reaction be detectedelectrochemically. These oxidoreductase enzyme systems include, forexample: 1) the secondary alcohol dehydrogenase (S-ADH)-catalyzedreduction of acetone with concomitant NADPH consumption; 2)S-ADH-catalyzed reduction of acetone with concomitant NADH consumption;3) acetone carboxylase reaction coupled to β-hydroxybutyratedehydrogenase consumption of NADPH; 4) acetone carboxylase reactioncoupled to β-hydroxybutyrate dehydrogenase consumption of NADH; 5)acetone carboxylase reaction ATP hydrolysis coupled to NADPHconsumption; 6) acetone carboxylase reaction ATP hydrolysis coupled toNADH consumption; 7) S-ADH reaction NADP⁺ formation coupled to H₂O₂formation; 8) S-ADH reaction NAD⁺ formation coupled to H₂O₂ formation;9) acetone carboxylase reaction ATP hydrolysis coupled to H₂O₂formation; 10) acetone carboxylase reaction coupled to β-hydroxybutyratedehydrogenase NADP⁺ formation coupled to H₂O₂ formation; 11) acetonecarboxylase reaction coupled to β-hydroxybutyrate dehydrogenase NAD⁺formation coupled to H₂O₂ formation; 12), acetone mono-oxygenase coupledto NADPH oxidation; 13) acetone mono-oxygenase coupled to NADHoxidation; 14) acetone mono-oxygenase coupled to H₂O₂ formation; and 15)acetone monooxygenase-catalyzed NAD(P)+ formation coupled to H₂O₂formation.

In all of these enzyme systems, the pyridine nucleotide or hydrogenperoxide is detectable electrochemically, though other detection meansknown in the art can be utilized.

The use of enzymes as bioactive interfaces is well known in the art, andsuch interfaces are used in analytical methods of detecting electronictransduction of enzyme-substrate reactions. Direct electrical activationof enzymes such as redox enzymes permits stimulation ofbioelectrocatalyzed oxidation or reduction of enzyme substrates. Rapidtransfer of electrons between an electrode and a given redox enzymeresults in current generation corresponding to the rate of turnover ofthe electron exchange between the substrate and biocatalyst. In otherwords, the transduced current of the system correlates with enzymesubstrate concentration. Electrical contacting of redox proteins in abiosensor and the electrode support contained therein may be mediated bydirect electron transfer with electrode surfaces. Redox enzymes lackingdirect electrical communication with electrodes may achieve electricalcontact by mediated electron transfer via active charge carriers. Anelectron relay may be oxidized or reduced at an electrode surface, anddiffusion of the oxidized or reduced relay into enzyme results in shortelectron transfer distances with respect to the active redox center formediated electron transfer and, thus, electrical activation of abiocatalyst.

Detection Means

The acetone-selective enzyme system, in acting upon the acetonesubstrate, generates an electrochemically or non-electrochemicallydetectable product or by-product directly, or the enzyme system willalso include at least one further component. The further component maybe: one or more additional enzyme(s) forming an enzymatic pathwayutilizing the product or by-product of the initial enzymatic acetonereaction to thereby generate a photometrically or electrochemicallydetectable product or by-product; or at least one signal mediator; orboth the additional enzyme(s) and the signal mediator(s). The signalmediator(s) may be selected from, for example: indicators, such as apH-change indicators; electron transfer mediators; photometricmediators, and other components.

In an electrochemical embodiment of the invention, an acetone-specificredox enzyme or enzyme system is selected that utilizes anelectrochemically detectable cofactor, such as NADH, or generates aby-product, such as H₂O₂, during the course of the enzymatic reaction.These enzyme systems can selectively detect acetone in biologicalsamples, such as breath or biological fluids. However, detection ofacetone is not limited to electrochemical means, and the enzyme systemof this invention may be used in other types of devices, for exampledevices employing known V, fluorescence, or other suitable methods ofdetecting acetone-specific enzyme-substrate interactions.

Non-Electrochemical Detection Means

Non-electrochemical detection involves, for example, any colorimetric orphotometric detection mode known in the art (for example, anycolorimetric, spectrometric, spectrophotometric, luminescence-based,chemiluminescence-based, or fluorescence-based detection method.)

A fluorescence detection device has the following minimum requirements:it must be light-tight to eliminate stray light from its surroundings,its fluors must be stored in the dark to prevent photobleaching (that isincrease shelf life), and its optics must be at a 90° angle. A diodeemitting the desired excitation wavelength can function as the lightsource, and a PMT can function as the detector. These need not beelaborate since both the excitation and emission max of the fluor areknown, and these are the only wavelengths required. The same breathcollection and acetone partitioning apparatus used in an enzymeelectrochemical device can be used in a fluorescence device. A portablefluorescence detector for aflatoxin has been described in the literature(M A Carlson et al., An automated handheld biosensor for aflatoxin,Biosens. Bioelectr. 14:841 (2000)), so a precedent for a portablefluorescence detector exists.

Both direct and indirect fluorescence allows the detection of acetonefrom both breath and body fluids. The acetone-specific enzymes and theircofactors can be immobilized on a disposable strip using conventionalentrapment techniques. When acetone diffuses through the immobilizationmedium to the enzyme, the acetone will be chemically altered.Unfortunately, acetone itself is not fluorescent and cannot bederivatized inside the detection device. Thus another reagent needs tobe derivatized with a fluorophore or a fluor needs to be added to thesystem to monitor the reaction. For the secondary alcohol dehydrogenasesystem, NADH consumption can be monitored, while the acetone carboxylasesystem can use ATP-analogs. As the NADH or ATP-analog is consumed,fluorescence intensity should decrease. Since the reaction with acetoneis stoichiometric, fluorescence intensity is proportional to acetoneconcentration. The H₂O₂-generating systems can use H₂O₂ and anadditional fluor. In these systems, H₂O₂ production causes an increasein fluorescence intensity that is proportional to acetone concentration.

We have verified that NADH in 100 mM phosphate buffer, pH 7.6, emitslight directly at approximately 470 nm when excited with 342 nm light;these data agree with those reported in the literature (M A Carlson etal., 2000). In addition, NADH direct fluorescence has a 0.1-10 μM linearworking range, is independent of pH from pH 6-13, decreases in intensity1.6% per ° C., and exhibits little altered fluorescence intensity in thepresence of cations and enzymes below pH 10 (See P W Carr & L D Bowers,Immobilized Enzymes in Analytical and Clinical Chemistry, In ChemicalAnalysis. A Series of Monographs on Analytical Chemistry and ItsApplications (P J Elving & J D Winefordner, eds.; vol. 56, p. 122(Wiley-Interscience, New York, 1980), and references contained therein).Several groups have described the use of direct NADH fluorescence tomonitor enzymatic activity (A K Williams & J T Hupp, Sol-gelencapsulated alcohol dehydrogenase as a versatile, environmentallystabilized sensor for alcohols and aldehydes, J. Am. Chem. Soc. 1998,120:4366; and V P Iordanov et al., Silicon thin-film UV filter for NADHfluorescence analysis, Sens. Actuat. A, 2002, 97-98:161).

Indirect fluorescence of NADH can be detected using the dye rhodamine123. Non-radiative energy transfer (also called fluorescence resonanceenergy transfer, FRET) occurs between the excited states of NADH andrhodamine 123. FRET is a well-known technique for determining theproximity of two species, i.e. FRET is utilized as a “molecularyardstick” both in vitro and in vivo. In this context of anacetone-specific enzyme system, a donor fluorophore, e.g., NADH,transfers its excited state energies to the acceptor fluorophore,rhodamine 123. (R P Haugland, Handbook of Fluorescent Probes andResearch Products, 2002 (9.sup.th ed.; Molecular Probes, Inc.; Eugene,Oreg.); K Van Dyke et al., eds. Luminescence Biotechnology. Instrumentsand Applications, 2002 (CRC Press; Boca Raton, Fla.) and referencescontained therein). The NADH-rhodamine 123 FRET method has beensuccessfully employed in other enzymatic assays (M H Gschwend et al.,Optical detection of mitochondrial NADH content in intact humanmyotubes, Cell. Mol. Biol. 47:OL95 (2001); H. Schneckenberger et al.,Time-gated microscopic imaging and spectroscopy in medical diagnosis andphotobiology, Opt. Eng. 33:2600 (1994)). Bioluminescence resonanceenergy transfer, or BRET, may also be used in conjunction with anacetone-specific enzyme system according to the present invention. InBRET, the donor fluorophore is replaced by a luciferase. Bioluminescencefrom luciferase in the presence of a substrate excites the acceptorfluorophore. BRET has also been applied in vitro and in vivo (K Van Dykeet al., 2002).

ATP can be derivatized with a fluorophore for indirect fluorescence.Several commercially available dyes include BODIPY ATP andtrinitrophenyl ATP (Haugland, 2002). These analogs change theirfluorescence intensity or become fluorescent when bound to an enzyme'sATP binding site.

Indirect fluorescence detection of H₂O₂ has also been reported (Carr &Bowers, 1980). These methods utilize dyes that reduce the peroxide toH₂O and are themselves oxidized. Homovanillic acid(4-hydroxy-3-phenylacetic acid) and p-hydroxyphenylacetic acid are amongthe most commonly used in clinical chemistry (Carr and Bowers, 1980). Acommercially available kit uses the dye Amplex Red for fluorescencedetection of H₂O₂ (Haugland, 2002).

Any fluorescent dyes and fluorescence-detectable enzyme substrate orcofactor analogs can be used in a fluorescence device to detect acetonein breath or bodily fluids.

Chemiluminescence (CL) and electrogenerated chemiluminescence (ECL)(collectively referred to herein as “(E)CL”) are widely used in medicaldiagnostics and analytical chemistry (C Dodeigne et al.,Chemiluminescence as a diagnostic tool: A review, Talanta 2000, 51:415;K A Fhnrich et al., Recent applications of electrogeneratedchemiluminescence in chemical analysis, Talanta 2001, 54:531).Enzyme-based (E)CL systems are sensitive and specific, and many CLsystems are used with enzyme cycling to detect H₂O₂ (Dodeigne et al.,2000). (E)CL can detect picomolar (pM; 10⁻¹² M) concentrations ofanalyte over a wide linear range (Dodeigne et al., 2000; Fhnrich et al.,2001). An (E)CL device can be constructed in accordance with thefollowing principles. Since the reaction itself emits light, an (E)CLdevice does not need a light source. A photomultiplier tube (PMT) canfunction as the detector; (E)CL is visible to the unaided, dark-adaptedeye. A battery can be the power source for ECL. ECL requires electrodesand a source of applied potential. Like a fluorescence detection device,(E)CL devices need to be light tight and their reagents need to beprotected from light until use. Also like fluorescence, (E)CL requiresderivatized reagents or additional enzymes and reagents to detectacetone. (E)CL devices can be used with disposable strips (B D Leca etal., Screen-printed electrodes as disposable or reusable optical devicesfor luminol electrochemiluminescence, Sens. Actuat B. 2001, 74: 190) andcan be miniaturized (Y Lv et al., Chemiluminescence biosensor chip basedon a microreactor using carrier airflow for determination of uric acidin human serum, Analyst 2002, 127:1176). An optical electrode (oroptrode) can be fabricated using an acetone-specific enzyme systemaccording to the present invention. For example, an optrode such as thatused in a glucose optrode that uses ECL, may be employed (see CH Wang etal., Co-immobilization of polymeric luminol, iron(II)tris(5-aminophenanthroline) and glucose oxidase at an electrode surface,and its application as a glucose optrode, Analyst 2002,127:1507)).

The most common CL systems involve the detection of H₂O₂ or anotherreactive oxygen species (Carr & Bowers, 1980; Haugland, 2002; Dodeigneet al., 2000; K Van Dyke et al., 2002) and references containedtherein). The classic system is luminol-peroxidase. In basic solution,H₂O₂ oxidizes luminol to an excited amino-phthalate ion; the excitedamino-phthalate ion emits a 425-nm photon to return to its ground state.When used in medical diagnostics, this reaction is catalyzed withhorseradish peroxidase (HRP) (Carr & Bowers, 1980; Dodeigne et al.,2000). Thus any enzyme system that produces H₂O₂ or requires a cofactorthat can react with additional reagents to form H₂O₂ can be used in a CLdevice. The H₂O₂-generating systems described herein can use luminol-HRPdirectly for acetone detection. These enzyme cycling schemes increasethe light emission over time because the substrates are continuouslyrecycled (Dodeigne et al., 2000). While luminol itself is frequentlyused in CL, its improved analogs can also be used in a CL-based detectoraccording to the present invention, in place of luminol, in order toincrease the sensitivity. Examples of such analogs are those describedin Carr & Bowers, 1980; and Dodeigne et al., 2000.

NADH detection using CL is a common technique (Dodeigne et al., 2000).For example, in the presence of 1-methoxy-5-methylphenaziniummethylsulfate, NADH reduces O₂ to H₂O₂ which generates light using theluminol-peroxidase system (Dodeigne et al., 2000). For an acetonemonitor, the O₂ in ambient air is sufficient to detect acetone usingthis system. NADH also reacts with oxidized methylene blue to form H₂O₂that reacts with luminol (Carr and Bowers, 1980). NADH can also act as aCL quencher. The fluorescence intensity of the substrate ALPDO isdecreased in the presence of NADH and HRP (Van Dyke et al., 2002). NADHalso can be used with Ru(bpy)₃ ²⁺ for ECL (E S Jin et al., Anelectrogenerated chemiluminescence imaging fiber electrode chemicalsensor for NADH, Electroanal, 2001, 13(15):1287). Rhodamine Bisothiocyanate can also be used for ECL detection of H₂O₂ (Fhnrich etal., 2001). ECL also offers another advantage in that, by use of aproperly poised electrode, the electroactive species can be regeneratedat the electrode surface. Regeneration both conserves reagents andallows durable and/or “reagentless’ sensors. All these systems can beused in a (E)CL device interfaced to an acetone-specific enzyme systemaccording to the present invention.

CL is widely used to quantitate ATP simply and sensitively (Carr &Bowers, 1980). The enzyme luciferase catalyzes the reaction of ATP andluciferin to produce excited-state oxyluciferin, which returns to itsground state with the emission of a 562-nm photon (Carr & Bowers, 1980;Haugland, 2002). The quantum yield for this reaction is very high; 10⁻¹⁴mol ATP can be detected. A kit for this reaction is commerciallyavailable (Haugland, 2002). Because luciferase is the enzyme that causesfireflies to “glow,” this reaction is referred to as bioluminescence.Both native and recombinant luciferase are commercially available, andseveral groups have reported using bioluminescence ATP assays toquantify biological analytes (P Willemsen et al., Use of specificbioluminescence cell lines for the detection of steroid hormone[ant]agonists in meat producing animals, Anal. Chim. Acta 2002, 473:119;S J Dexter et al., Development of a bioluminescent ATP assay to quantifymammalian and bacterial cell number from a mixed population, Biomat.2003, 24:nb27). In addition to the luminol-HRP system, H₂O₂ can also bedetected using peroxyoxalic acid derivatives (Dodeigne et al., 2000).H₂O₂ can also be detected with CL non-enzymatically with ferricyanide asthe catalyst (Dodeigne et al., 2000). In these (E)CL systems, theacetone-specific enzymes described herein either produce H₂O₂ or requirecofactors that can be utilized to form H₂O₂.

Optical biosensors use photometric detection (that is, absorbance,fluorescence) of substrates consumed or products formed by the reactioncatalyzed by the enzyme system incorporated into the sensor. Theacetone-specific enzyme reactions described may be monitored by severalphotometric methods-namely by measuring NAD(P)H absorbance at 340 nm forthe pyridine nucleotide-dependent enzymes or absorbance of thequinoneimine dye for the H₂O₂ forming enzyme systems. For the later,addition of a peroxidase allows detection of H₂O₂ by catalyzing thereduction of H₂O₂ with concomitant oxidation of a dye compound that uponoxidation absorbs at a specified wavelength. Peroxidase enzymes (forexample, commercially available horseradish peroxidase) typically havebroad substrate specificities so several different electron donorcompounds may be used. NAD(P)H consumption may also be measured byfluorescence detection (excitation at 350 nm and emission at 450 nm).

Calorimetry may be employed as a detection means in an acetone-specificsensor according to the present invention. Chemical reactions aretypically either exo- or endothermic; that is, they release or absorbheat as they occur. Calorimeters detect and measure this heat bymeasuring a change in the temperature of the reaction medium (KRamanathan & B Danielsson, Principles and applications of thermalbiosensors, Biosens Bioelectr. 16:417 (2001); B Danielsson, EnzymeThermistor Devices. In Biosensor Principles and Applications. Vol. 15,pp. 83-105 (L J Blum & P R Coulet, eds.; Bioprocess Technology Series,volume 15; Marcel Dekker, Inc: New York, 1991, pp. 83-105, andreferences contained therein). Thus, the action of an acetone-specificenzyme or enzyme system may be monitored calorimetrically. Calorimetershave been designed that are sensitive enough to detect proteinconformational changes, and calorimetry has been used to study manyenzymatic reactions in detail (M. J. Todd & J Gomez, Enzyme kineticsdetermined using calorimetry: a general assay for enzyme activity? Anal.Biochem. 2001, 296:179 (2001)).

The major advantage of calorimetry is the lack of derivatizationrequired for analysis (Danielsson, 1991). Since most reactions involveheat exchange, and this heat is detected, no chromophores, fluorophores,luminophores, “mediators,” or other modifications of the analyte arerequired. Reagents and analytes can be used “as is.” This allows theanalysis of both reactions that lack a chromophore or fluorophore and/orwould be difficult or impossible to derivatize or couple to thegeneration of an electroactive species.

Miniaturized or chip-based thermosensors have been reported in theliterature (Ramanathan & Danielsson, 2001; B Xie & B Danielsson,Development of a thermal micro-biosensor fabricated on a silicon chip.Sens. Actuat. B 6:127 (1992); P Bataillard et al., An integrated siliconthermopile as biosensor for the thermal monitoring of glucose, urea, andpenicillin. Biosen. Bioelect. 8:89 (1993)). These devices range fromradically arranged thermopiles on freestanding membranes to groups ofthermopiles constructed on silicon/glass microchannels. These deviceshave been used to detect specific, single enzymatic reactions(Danielsson, 1991; Xie & Danielsson, 1992; Bataillard et al., 1993).Moreover, two groups have reported thermosensors for glucose (B Xie etal., Fast determination of whole blood glucose with a colorimetricmicro-biosensor, Sens. Actuat. B 15-16:141 (1993); M J Muehlbauer etal., Model for a thermoelectric enzyme glucose sensor, Anal. Chem. 61:77(1989); B C Towe & E J Guilbeau, Designing Medical Devices, 1998,http://lsvl.la.asu.edu/asubiotech/slideshow/slide19.html (accessedJanuary 2002). Preliminary experiments using a conventional calorimeterindicate that the secondary alcohol dehydrogenase-acetone reaction isexothermic (data not shown).

For the acetone monitor described herein, the acetone-specific enzymesand their cofactors can be immobilized on a thermopile via conventionalentrapment methods. The enzymes and reagents associated with the coupledelectrochemical detection, electrochemical mediators, and “photonic”mediators (luminophores) are unnecessary for calorimetry. The reactioninvolving acetone can be monitored directly without modification orderivatization. When acetone in the breath or fluid sample diffusesthrough the immobilization medium and encounters the enzyme, the acetonewill be chemically altered. This reaction will generate or absorb heat,causing a temperature change. Comparison of this temperature with thatof a reference thermopile will quantify this heat; the measurement isdifferential. The quantity of heat released or absorbed is proportionalto the analyte concentration.

For breath collection, partitioning the acetone from the gas phase tothe liquid phase, that is, condensation, is exothermic. The reference ordual thermopile can compensate for this heat. Thus an enzymecolorimetric acetone monitor can use the same breath collectionapparatus as an enzyme electrochemical acetone monitor except for theaddition of the dual thermopile.

The entire enzyme colorimetric device needs to be sufficiently insulatedto prevent heat exchange with its surroundings. Except forelectrochemical detection, other aspects of the device, such as enzymestability, specificity, device portability, etc., described in thisdocument are the same as those for an enzyme electrochemical device.

Thus, useful methods for achieving signal transduction in biosensorsaccording to the present invention include not only electrochemical(amperometric or potentiometric), but also optical or photometric(including colorimetry, fluorescence-based techniques, orchemiluminescence-based techniques), and colorimetric means, all ofwhich are useful in application to acetone biosensor signaltransduction.

Therefore, although electrochemical detection means are described andexemplified in detail herein, the enzyme systems of the invention arenot limited to use in biosensors employing electrochemical detectingstrategies. Other detection strategies may be suitably integrated into abiosensor specific for acetone in biological samples. Photometricassays, such as assays in which changes in the amount of light absorbedin a reaction solution over time may be used. Likewise, assays in whichchanges in fluorescence or changes in sample turbidity may be employedfor detecting acetone-specific enzyme-substrate interactions. Suchphotometric assays are discussed hereinbelow. Redox potentials of H₂O₂and colorimetric/photometric detection of coenzymes is discussed byBergmeyer. Photometric assays for enzymatic activity are generallydescribed by John in “Photometric Assays”. An NADH-consumption measuringelectrode is disclosed by Hart et al. in a 1999 article published inElectroanalysis. Vanysek discloses redox potentials in general, andoxidation-reduction potentials of various compounds suitable for use inbiochemical applications are disclosed by Voet & Voet.

An enzyme system employing S-ADH coupled to alanine dehydrogenase wassuccessfully monitored spectrophotometrically for NADH formation. Inaddition, since the reaction also generates ammonium ion, an opticalsensor for NH₄ ⁺ can be employed as the detection means for such anenzyme system. One such optical means is described by TD Rhines and M AArnold, Fiber-optic biosensor for urea based on sensing of ammonia gas,Anal Chim. Acta, 1989, 227:387; several enzyme-based amperometric NH₄ ⁺sensors are commercially available. For acetone detection, ammoniaproduction can be coupled to the secondary alcohol dehydrogenase systemas above; ammonia concentration would then be proportional to acetoneconcentration. Another enzyme scheme to couple acetone to ammoniaproduction is the following:Acetone+NADH+H⁺→2-propanol+NAD⁺ (S-ADH)glutamate+NAD⁺+H₂O-ketoglutarate+NH₄ ⁺+NADH (glutamate dehydrogenase)

This second scheme can be used either optically or amperometrically todetect acetone. Additionally, the NADH is recycled. Likewise, an enzymesystem in which acetone carboxylase is coupled to glutamatedehydrogenase, generates NH₄ ⁺ and so can be detected optically oramperometrically and correlated with acetone concentration.

Electrochemical Detection Means

Amperometric biosensors work by generating current between twoelectrodes by enzymatically producing or consuming a redox-activecompound. Several examples of amperometric acetone biosensor schemeshave been described in which NAD(P)H or H₂O₂ are consumed or generatedenzymatically in response to the presence of acetone. In examples wherethe transducer is H₂O₂, an alternative means to monitor the reactionamperometrically could be to employ a Clark-type oxygen electrode andmeasure a decrease in O₂ concentration. For example, in the case for thesecondary alcohol dehydrogenase (S-ADH) coupled to H₂O₂ formation, theenzyme system catalyzes the following:acetone+NADH+H⁺→2-propanol+NAD⁺ (S-ADH)lactate+NAD⁺→pyruvate+NADH+H⁺ (lactate de-hydrogenase)pyruvate+Pi+O₂→acetylphosphate+CO₂+H₂O₂ (py-ruvate oxidase)

Oxygen is then reduced/consumed at the cathode generating aconcentration gradient between the electrode and the bulk solution. Therate of electrochemical reaction is dependent on the oxygenconcentration in solution.

Potentiometric biosensors employ ion-selective electrodes in which therelease or consumption of ions during an enzyme reaction is measured(for example, H⁺, CN⁻, NH₄ ⁺) (1, 2, 3). For example, a potentiometricbiosensor for measuring acetone concentration can be utilized where NH₄⁺ formation is coupled to the reaction catalyzed by S-ADH and alaninedehydrogenase as follows:acetone+NADH+H⁺→2-propanol+NAD⁺ (S-ADH)alanine+NAD⁺→pyruvate+NADH+H⁺+NH₄ ⁺ (alanine dehydrogenase)

(Photometric data for this system has already been obtained to verifyits utility for acetone-specific signal transduction: see the discussionunder the “Results” section, below).

A very similar system can be utilized with acetone carboxylase(“β-OH-butyrate dehydr.” being “β-hydroxbutyrate dehydrogenase”):acetone+ATP+CO₂→acetoacetate+AMP+2P_(i) (acetone carboxylase)acetoacetate+NADH+H⁺→β.-hydroxybutyrate+NAD⁺ (β-OH-butyrate dehydr.)alanine+NAD⁺→pyruvate+NADH+H⁺+NH₄ ⁺ (alanine dehydrogenase)

Another type of electrochemical biosensor that may be employed is alight-addressable potentiometric sensor. In one embodiment of such adevice, the acetone-specific enzyme system(s) may be applied to (e.g.,immobilized to the surface of) a potentiometric sensing means such asthat described, for sensing glucose, in A Seki et al., Biosensors basedon light-addressable potentiometric sensors for urea, penicillin, andglucose, Anal. Chim. Acta 373(1):9-13 (2 Nov. 1998).

In designing an acetone-specific biosensor according to the invention,various enzymatic by-products and/or factors may be employed for theproduction of electrochemical signals. One group includes organiccofactors, such as NAD, NADH, NADP, NADPH, FAD, FADH, FMN, FMNH,Coenzyme A, Coenzyme Q, TTQ (Tryptophan Tryptophylquinone) and PQQ(Pyrroloquinolinequinone). For example, a PQQ-dependent dehydrogenasemay oxidize isopropanol. Electrons from this reaction may be transferredthrough PQQ, which is reduced, and can be oxidized at the electrode orwith an intervening enzyme. Other vitamins may also be used.

Enzymatic reaction by-products useful in the invention include hydrogenperoxide and ammonium.

Energetic molecules may also be used in the invention for couplingacetone metabolism to electrochemically measurable signals, including:ATP, ADP, AMP, GTP, GDP and GMP. Neither these molecules nor phosphatecan be detected directly, but can be detected through coupling to aredox-by-product-producing enzyme system.

These by-products, cofactors, and energetic molecules can also bedetected by non-electrochemical means as described above.

Signal Mediators

Electron transfer mediators are redox-reversible species that may beused to transfer electrons between (that is to or from) the electricallypotentiated surface of an electrode and an organic species (such as aco-factor) involved in an enzymatically catalyzed reaction. Examples ofelectron transfer mediators include: ferrocene and derivatives,ferricyanide, hydroquinone, benzoquinone and derivatives,2,6-dichloroindophenol, methylene blue, phenylenediamine andderivatives, phenoxazine and derivatives (for example, Meldola's blue,that is 8-dimethylamino-2,3-benzophenoxazine), and phenazinealkosulfates (for example, phenazine methosulfate, phenazineethosulfate). In a given embodiment, one or more than one species ofelectron transfer mediator may be used.

Electron transfer mediators can be used to improve the kinetics ofelectron transfer in a given enzyme-coupled electrode system, sinceorganic cofactors may easily impair detector functions. This impairmentis caused by the creation of free radicals via singly transferringmultiple electrons between organic species and the electricallypotentiated surface of the electrode. These free radicals then canexhibit dimer and/or polymer formation at the electrically potentiatedsurface, which fouls the surface of the electrode, thereby inhibitingefficient electron transfer. Electron transfer mediators can be employedto avoid this fouling of electrodes. Electron transfer mediators mayalso be used in situations where a shift in electrode voltage isdesired, for example, where the preferred voltage for use in thereaction system without such a mediator happens to be a potential atwhich too much electrical interference (“noise”) occurs. An electrontransfer mediator may be added in order to permit a shift in the appliedvoltage to a different voltage region in which less noise occurs.Examples of diffusional electron-transfer mediators applicable toimmobilized enzymes such as glucose oxidase, horseradish peroxidase, andthe like, are set forth in Table 5 of Willner and Katz.

Preferred mediators useful in multi-electron transfers for reduced formsof, for example, NADH, NADPH, FADH, FMNH, Co-Q, PQQ, include, forexample: ferrocene and derivatives, ferricyanide, hydroquinone,benzoquinone and derivatives, 2,6-dichloroindophenol, methylene blue,phenylenediamine and derivatives, phenoxazine and derivatives (forexample, Meldola's blue, that is 8-dimethylamino-2,3-benzophenoxazine),and phenazine alkosulfates (for example, phenazine methosulfate,phenazine ethosulfate).

A second group of mediator factors that may be employed for theproduction of electrochemical signals include inorganic cofactors suchas Pt, Os, V, Mn, Fe, Co, Ni, Cu, Mo, and W (see Holm et al., Aspects ofMetal Sites in Biology, Chem. Rev. 1996. 96, p. 2239-2314). Some usefulenzymes contain a heme center, and thus iron (for example, cytochromeP450 monooxygenase). Also useful is amine oxidase, which contains Cu. Inan alternative embodiment, a “photometric mediator” may be added to theenzyme system in order to react with a product or by-product of theenzymatic reaction(s) and thereby generate a derivative that can be, forexample, photochemically, calorimetrically, fluorometrically, or (UV orIR) spectrometrically detected. Thus, the addition of such a“photometric mediator” may be characterized as permitting the conversionof a result of the enzymatic reaction, that is a product or by-product,into a photometric signal. For example, in the case of enzymaticallycatalyzed redox reactions, a chromogenic redox indicator such as, forexample, a tetrazolium salt, may be used as the photometric mediator.Many such chromogenic redox indicators are known in the art. Examples oftetrazolium salts include, but are not limited to:3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-2H-tetrazolium bromide (MTTbromide);(3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium,inner salt (MTS; available from Promega Corp., Madison, Wis.); and(5-cyano-2,3-ditolyl tetrazolium chloride) (CTC). Such photometricmediators can be used, for example, to convert the redox “signal” of anelectron transfer mediator into a photometrically detectable signal.

The enzymes and other components of the enzyme system may be immobilizedin a gel layer disposed upon the electrode surface. Any of the variousgels known in the art as useful for immobilization of biologics in thepresence of an electrode may be used. For example, a method such as isdescribed in PCT/US02/16140 (filed May 21, 2002) may be used toimmobilize the biologic components of an acetone-specific enzyme systemin a polyurethane hydrogel disposed upon an electrode.

Enzyme Arrays and Multi-Enzyme Systems

In addition to monitoring acetone in a sample per se, acetone-specificenzyme systems may be useful in biosensors having more than one enzymesystem for detecting multiple substrates in a given biological sample.For example, an electrode linked to an acetone-specific enzyme systemmay enable subtraction of an acetone signal from an ethanol detector.Such a set up could be configured in an array, wherein at least twodifferent detection modes or at least two different detectors would beoperative for detecting ethanol and acetone. Such an array would beuseful to correct for acetone interference in ethanol breathalyzeranalyses.

Fluorescence detection can also be accomplished using arrays.Fluorescence sensor arrays have been described in the literature. Theyhave been used for such complex samples as wine aromas, perfume, andgenes. Fluorescence sensor arrays employ fluorescent or chromogenic dyesor substrates that covalently attached to polystyrene beads in wells onthe distal face of an optical fiber (D. R. Walt, Imaging optical sensorarrays. Curr. Opin. Chem. Biol. 6:689 (2002)). A high-density opticalarray can contain several types of dyes or substrates for differentanalytes. The array is exposed to each possible component individually,then to the sample. Pattern recognition is employed to deduce thecomposition of the sample. In the case of breath or bodily fluidcomponents, the acetone-specific enzymes, cofactors, and chromogenic orfluorogenic dyes can be covalently attached to a portion of beads, whileenzymes specific for other analytes, such as ethanol, can be attached toother beads. Each bead will “light up” upon exposure to its targetanalyte.

An enzyme-based fluorescence or chromogenic array has never been appliedto the detection of acetone.

Uses for an Acetone-Specific Enzymatic Biosensor

Breath acetone monitoring is a useful tool for monitoring effectivenessand compliance of subjects on weight loss diets. Ketosis can bemanipulated by exercise and dieting choices, even between two diets withequal energy balance. The response time for reflecting diet and exercisechoices in breath acetone levels is in the order of 2-3 hours, and was abetter indicator of fat loss than urine ketone analysis.

A home acetone diagnostic biosensor would be useful in aiding subjectmanagement of Type 1 and Type 2 diabetes. Such biosensors would enablesubjects to monitor weight loss, to detect signs of the onset ofketoacidosis, and to control sugar intake with respect to insulinavailability, especially in Type 1 diabetics. Indicators suggest thatweight loss success would be improved if subjects could share dailyacetone measurements with health care professionals and peers via theInternet and weekly support group meetings. Use of the inventiveacetone-specific detection system is not limited to management ofobesity and diabetes. It is contemplated that the acetone-specificbiosensors described herein would be useful for managing any disease inwhich acetone production is an indicator of pathology.

In addition, the acetone-specific enzyme system may prove to be a highlyeffective means of monitoring subject compliance with prescribedtherapeutic regimes via drug tagging with acetoacetate or a derivativethereof. The degradation of acetoacetate to acetone could be measuredvia a biosensor containing the inventive acetone-specific enzyme system,thereby improving the ability of health care professionals to track thedosing and bioavailability of the corresponding tagged drug.

EXAMPLES

Acetone-specific enzyme systems and acetone sensors utilizing thesesystems have been developed. Enzyme identification and/or purification,enzyme characterization and selection, enzyme-plus-cofactor systems,multiple-enzyme-plus cofactor systems, coupled enzyme systems providinglinear (stoichiometric) acetone detection, coupled enzyme systemsproviding amplified (for example, exponential) acetone detection, enzymeand enzyme system stability testing, acetone vapor-to-liquidpartitioning studies, and enzyme-mediated acetone sensor devices (bothelectrochemical and non-electrochemical devices) that utilize suchsystems sensors are disclosed below in particular exemplifiedembodiments. These examples are provided for exemplification and are notintended to limit the invention. Particular embodiments employingacetone-specific enzyme systems in enzyme-based electrochemical andnon-electrochemical sensors is described below.

Materials & Methods

Materials. Acetone carboxylase from X. autotrophicus strain Py2 andisopropanol-grown X. autotrophicus strain Py2 cell paste were obtainedfrom Professor Scott A. Ensign at Utah State University, Logan, Utah.Acetone carboxylase from Rhodobacter capsulatus B10 (ATCC 33303),acetone-grown R. capsulatus B10 cell paste, propane-grown Mycobacteriumvaccae JOB5 (ATCC 29678) cell-free extracts, and propane-grownRhodococcus rhodochrous B276 (ATCC 31338) cell paste were also obtainedfrom Professor Ensign, and all of these bacterial strains are publiclyavailable. Secondary alcohol dehydrogenase from X. autotrophicus strainPy2 and isopropanol-grown X. autotrophicus strain Py2 were isolated fromcell paste; and exemplary, publicly available secondary alcoholdehydrogenases are described in Table 1. Pyruvate kinase (EC 2.7.1.40),myokinase (EC 2.7.4.3), pyruvate oxidase (EC 1.2.3.3), horseradishperoxidase (EC 1.11.1.7), lactate dehydrogenase (EC 1.1.1.28), malicenzyme (EC 1.1.1.40), alcohol dehydrogenase (EC 1.1.1.2), alaninedehydrogenase (EC 1.4.1.1), alcohol dehydrogenase (EC 1.1.1), alcoholoxidase (EC 1.1.3.13), and .beta.-hydroxbutyrate dehydrogenase (EC1.1.1.30) were purchased from Sigma (St. Louis, Mo.). All otherchemicals and reagents used were analytical grade. All solutions wereprepared in 18 M water (Millipore). TABLE 1 Information for SomePublicly Available S-ADH Enzymes Cofactor Organism Source Reference(s)[& Comments] NADPH Thermoanaerobium Sigma Chem. Co. R J Lamed et al.,Enzyme & Microb. brockii (catalog no. Technol., 3: 144 (1981); A8435) RJ Lamed & J G Zeikus, Biochem. J, 195(1): 183-90 (Apr. 1, 1981); ABen-Bassat et al., J Bact., 146(1): 192-99 (April 1981); [DNA sequenceavailable in GenBank (Acc. No. X64841)] NADH Mycobacterium ATCC 29678 JP Coleman et al., J Gen. Microbiol., vaccae strain 131(11): 2901-07(November 1985); JOB5 (Gram-positive) [Describes enzyme purification]NADH Pseudomonas ATCC 21439 C T Hou et al., Eur. J Biochem., sp. 6307[CRL 75] 119(2): 359-64 (October 1981); (Gram-negative) [Describesenzyme purification] NADH Xanthobacter ATCC PTA-4779 [Enzymepurification and autotrophicus characterization described herein] strainPy2 NADPH Thermoanaerobacter ATCC 33223 D S Burdette et al., Biochem. Jethanolicus 39E 316(1): 115-22 (May 1996); [Describes enzymepurification, gene cloning & DNA sequencing] NADH Candida utilis DSM70167; H Schutte et al., Biochim. et Biophys. (yeast) ATCC 26387 Acta,716(3): 298-307 (Jun. 16, 1982); [Describes screening for S-ADH activityin several yeast strains] NADPH “Anaerobic extremely Biocatalysts Ltd. —thermophilic bacterium” (Wales; catalog no. S300) NADH Candida boidinniFluka (Milwaukee, — (yeast) WI; catalog no. 91031) NADH Candida sp.NovaBiotec Dr. — (yeast) Fechter GmbH (Berlin, Germany; catalog no.“Isopropanol dehydrogenase (E.C. 1.1.1.80)”)

Enrichment and isolation of acetone-, isopropanol-, andpropane-utilizing microorganisms. Enrichment cultures were set up in 160mL serum bottles that were crimp-sealed with butyl rubber stoppers. Thebottles contained 10 mL mineral salts medium containing (in g/L):NaNH₄HPO₄ (1.74); NaH₂PO₄.xH₂O (0.54); KCl (0.04); MgSO₄x7 H₂O (0.2) and1 mL/L of a trace element stock solution (stock solution (in g/L):FeCl₂x4 H₂O (5.4); MnCl₂x4 H₂O (1.0); ZnSO₄x7 H₂O (1.45); CuSO₄x5 H₂0(0.25); concentrated HCl (13 mL/L); (NH₄)₆Mo₇O₂₄x4 H₂O (0.1); H₃BO₃(0.1); CoCl₂x6 H₂O (0.19)). The pH of the medium was adjusted to pH 7.2.Enrichments for propane-utilizing microorganisms were inoculated withabout 0.5 g of soil that had been purchased from a local supplier of topsoils, or with about 0.5 g of non-sterilized potting soil or organiccompost that had been purchased from a local supermarket.

Gaseous propane was added with a syringe to a 20% (v/v) concentration inthe headspace of the serum bottle. Enrichments for acetone- andisopropanol-utilizing microorganisms were set up in a similar way exceptthat substrates were added from a 1 M stock solution to a finalconcentration of 25 mM acetone, or 10 mM isopropanol. The enrichmentcultures were incubated on a shaker at 28° C. For the isolation ofsingle colonies, enrichment cultures were cultivated on mineral saltsmedium (as described above) containing 1.5% w/v agar (hereinafter“mineral salts agar”).

In a different set-up, enrichment cultures were started foracetone-utilizing microorganisms that could grow in the presence of aCO₂-trap. 20 mL of mineral salts medium with trace elements (see above)was filled into 250 mL baffled Erlenmeyer flasks. The medium wasinoculated with about 0.5 g of soil sample (see above). The Erlenmeyerflask was closed with a rubber stopper that had been modified to hold aglass bulb. The glass bulb contained about 0.5 mL of 50% (w/v) KOH. TheKOH trapped the CO₂ from the Erlenmeyer flask headspace. These set-upswere designed to enrich for acetone-utilizing microorganisms with anacetone carboxylase-independent pathway. The enrichment cultures wereincubated on a shaker at 28° C.

Enrichment cultures were transferred two to three times after turbidityindicated bacterial growth (usually after 3 to 5 days). To isolatesingle colonies, enrichment cultures were spread on mineral salts agarplates. For the isolation of propane-utilizers, the agar plates wereplaced in a 3.5 L anaerobic jar. Propane was added to the jar until apositive pressure of 0.3-0.5 bar was reached inside the jar. The jar wasplaced into an incubator at 28° C. For the isolation ofacetone-utilizing microorganisms, the agar plates were placed into a 1.4L desiccator. The desiccator contained two open glass vials with 3-4 mLneat acetone each. The desiccator was sealed with several layers ofPARAFILM (a wax-based sealing film, from American National Can, Chicago,Ill.) before it was placed in an incubator at 28° C. For the isolationof acetone-utilizing microorganisms that would grow in the presence of aCO₂-trap, agar plates were placed in a desiccator as described above. Inaddition to a vial with acetone, a vial containing 50% KOH (about 4 mL)was placed into the desiccator. Alternatively, for the isolation ofacetone- and isopropanol-utilizers, enrichment cultures were transferredto agar plates containing mineral salts medium plus acetone orisopropanol. Additional acetone or isopropanol was added onto a smallfoam plug that was placed inside the Petri dish. The Petri dish wassealed with several layers of parafilm to reduce evaporation ofsubstrates during incubation.

Colonies were visible on the agar plates after 5-10 days. Isolates weretransferred to fresh agar plates and incubated as described above.Isolated strains were also streaked onto nutrient agar to check forpurity. After several transfers on agar plates, 31 strains were isolatedthat looked different as evaluated by colony morphology. Eight strainswere isolated from propane enrichments (these were designated TDCC Prop1-8), eight strains were isolated from isopropanol enrichments (thesewere designated TDCC IP-1-8) and fifteen strains were isolated fromacetone enrichments (these were designated TDCC Ac 1-15). None of thesewere obtained from an acetone+KOH-trap enrichment.

Screening of isolates and culture collection strains for growth onacetone, propane, or isopropanol. Isolates and culture collectionstrains were screened for growth on acetone, propane, and isopropanol in60 mL-serum vials containing 5 mL of medium plus 0.005% (w/v) yeastextract as described above. The medium was inoculated from a singlecolony. Isopropanol was added from a stock solution to a finalconcentration of 8 mM. Cultures that showed more turbidity withsubstrates compared to cultures without substrates (medium blanks) wereconsidered hits. Hits were then screened for growth with acetone in thepresence of a CO₂-trap as described above.

Cultivation of isolates/strains, harvesting, and preparation ofcell-free extracts. Several isopropanol-utilizing strains werecultivated in larger batches for initial purification of secondaryalcohol dehydrogenase and experiments with cell-free extracts. StrainRhodococcus rhodochrous B276 (ATCC 31338) (formerly Nocardia corallinaB276) and strain TDCC IP-1 (and two additional strains: data not shown)were cultivated in 2×500 mL batches of mineral salts medium (forcomposition see above) plus 0.005% yeast extract. Isopropanol (8 mM) wasadded initially as carbon and energy sources. Cultures were incubated ona shaker (200 rpm) at 30° C. Growth was followed by monitoring theoptical density at 600 nm. More isopropanol was added at several timepoints when the growth rate decreased due to lack of substrate. A totalof about 96 mM isopropanol was added to the cultures. At the end of thelogarithmic growth phase, cells were precipitated by centrifugation (GSArotor, 8,500 rpm, 20 min.) at 4° C. The cells were washed once in 50 mMTris-HCl buffer, pH 7.5. The cell pellet was weighed and resuspended ina small volume of TRIS (2-amino-2hydroxymethyl-1,3-propane-diol) buffer(about 2 mL per g cells (wet weight)). Cells were frozen at −20° C.until further use. For the preparation of cell-free extracts, cells werethawed and broken by sonication (4×20 s, pulsed, 50% intensity). Celldebris and unbroken cells were precipitated by centrifugation for 5 min.at 14,000 rpm (in an Eppendorf benchtop centrifuge, Model 5417C,Brinkmann, Instruments, Inc., Westbury, N.Y.). Alternatively, for largerpreparations, the cell suspension was passed three times through amini-French pressure cell at 20,000 psi (137,895.2 kPa), and the lysatewas clarified by centrifugation at 6,000×g for 40 min at 4° C.

Protein purifications. Acetone carboxylase from X. autotrophicus strainPy2 and acetone carboxylase from R. capsulatus were purified asdescribed previously. Secondary alcohol dehydrogenase (S-ADH) from X.autotrophicus Py2 was purified via the following protocol. Cell-freeextracts (380 mL) of isopropanol-grown X. autotrophicus Py2 (150 g) wereprepared as described above, and applied to a 5×15 cm column ofDEAE-Sepharose FAST FLOW (Diethylaminoethyl cross-linked agarose beadmaterial; Catalog number 17-0709-10, Amersham Pharmacia Biotech,Piscataway, N.J.)) equilibrated in buffer A (25 mM MOPS(3-(N-morpholino)propanesulfonic acid), pH 7.6, 5% glycerol, 1 mMdithiothreitol) at a flow rate of 10 mL/min. After loading, the columnwas washed with 1000 mL buffer A and developed with a 2400 mL lineargradient of 90-290 mM KCl in buffer A. Fractions containing S-ADHactivity were pooled and dialyzed against 2 L of 25 mM potassiumphosphate (pH 6.2) containing 5% glycerol (buffer B) for 16 h at 4° C.The protein was then applied to a RED SEPHAROSE CL-6B (Procion Red HE-3Bdye-linked, cross-linked-agarose bead material, affinity matrix foraffinity chromatograph; Catalog number 17-0528-01, Amersham PharmaciaBiotech) column (1.5×10 cm) equilibrated in buffer B at a flow rate of 2mL/min. After washing the column with 30 mL of buffer B, S-ADH waseluted with 20 mL of buffer A containing 10 mM NAD⁺. Fractionscontaining S-ADH were then dialyzed against 2 L of buffer A for 16 h at4° C., concentrated by ultrafiltration (using a YM30 ultrafiltrationmembrane; catalog no. 13722, from Millipore, Bedford, Mass.), and frozenin liquid nitrogen. Partially purified S-ADH from bacterial screencultures was prepared as follows: cell-free extracts from 1 to 5 g ofcell paste were prepared as described above and applied to a 5 mL HITRAP Q column (quaternary, tetraethylammonium, cross-linked agarose beadmaterial for use as an anion exchange matrix; catalog number 17-1153-01,Amersham Pharmacia Biotech) equilibrated in 100 mM MOPS, pH 7.6,containing 5% (v/v) glycerol (buffer C). The column was washed with 10mL buffer C and developed with a 100 mL linear gradient of 0 to 100 mMNaCl in buffer C. Fractions containing S-ADH activity were pooled,concentrated to 0.5 mL using a 30 kDa MWCO (“molecular weight cut-off”)centrifugal membrane (catalog number UFV4BTK25, Millipore), and storedat −80° C.

Example 1

Acetone carboxylase coupled to NADH oxidation spectrophotometric assay.Assays were performed in 2 mL (1 cm path length) quartz cuvettes thathad been modified by fusing a serum bottle-style quartz top (7×13 mm atmouth), allowing the cuvettes to be sealed with a red rubber serumstopper. The reaction mix contained ATP (10 mM), MgCl₂ (11 mM),potassium acetate (80 mM), MOPS (100 mM), CO₂ (50 mM (1 mol CO₂(g) to 4mol potassium bicarbonate to maintain pH)), and 20 to 40 μg purifiedacetone carboxylase in a total volume of 1 mL at pH 7.6. The addition ofβ-hydroxybutyrate dehydrogenase (3 U) and NADH (0.2 mM) allowedacetoacetate formation to be coupled to the oxidation of NADH. Assayswere pre-incubated for 2 min. at 30° C. with all assay components exceptacetone. Assays were initiated by addition of acetone (2 mM). Thereaction was monitored by measuring the decrease in absorbance at 340 nm(ε₃₄₀ of 6.22 mM⁻¹cm⁻¹ for NADH) over time in an Agilent Technologies(Palo Alto, Calif.) model 8453 UV-Visible Spectroscopy System containinga thermostat-controlled cell holder at 30° C.

Acetone carboxylase coupled to H₂O₂ formation spectrophotometric assay.Assays were performed in 2 mL (1 cm path length) quartz cuvettes andcontained ATP (0.1 mM), MgSO₄ (10 mM), potassium acetate (80 mM),potassium phosphate (50 mM), CO₂ (50 mM (1 mol CO₂(g) to 4 mol potassiumbicarbonate to maintain pH)), 40 μg purified acetone carboxylase,phosphoenolpyruvate (2 mM), pyruvate kinase (20 U), myokinase (15 U),pyruvate oxidase (2 U), peroxidase (15 U), flavin adenine dinucleotide(0.01 mM), cocarboxylase (0.2 mM), 4-aminoantipyrine (0.5 mM), andN-ethyl-N-(2-hydroxy-3-sulfopropyl)-m-tol-uidine (0.02% w/v) in a totalvolume of 1 mL at pH 7.5. Coupling enzymes and reagents (that isphosphoenolpyruvate, pyruvate kinase, myokinase, pyruvate oxidase,flavin adenine dinucleotide, and cocarboxylase,) allowed ATP hydrolysisto be coupled to H₂O₂ formation (pyruvate oxidation). Addition ofperoxidase, 4-aminoantipyrine, andN-ethyl-N-(2-hydroxy-3-sulfopropyl)-m-toluidine allowed H₂O₂ formationto be monitored spectrophotometrically at 550 nm (6550 of 36.88 mM⁻¹cm⁻¹ for quinoneimine dye product) over time in a thermostat-controlledcell holder at 30° C. Assays were pre-incubated for 2 min. at 30° C.with all assay components except acetone. Assays were initiated byaddition of acetone (5 mM).

Example 2

Secondary alcohol dehydrogenase NADH oxidation spectrophotometric assay.Assays were performed in 2 mL quartz cuvettes and contained NAD(H) (0.2mM), potassium phosphate buffer (25 mM), and a source of enzyme(cell-free extracts, column fractions, or purified enzyme) in a totalreaction volume of 1 mL at pH 6.2 (for ketone reduction assays) or pH7.8 (for alcohol oxidation assays) at 30° C. Assays were pre-incubatedfor 1.5 min. at 30° C. with all assay components except substrate.Assays were initiated by addition of substrate (2.5 mM) and monitoredover time by measuring the change in absorbance at 340 nm (ε₃₄₀ of 6.22mM⁻¹ cm⁻¹ for NADH).

Secondary alcohol dehydrogenase coupled to H₂O₂ formationspectrophotometric assay. Assays were performed in 2 mL (1 cm pathlength) quartz cuvettes and contained potassium phosphate (50 mM), 1.5μg purified S-ADH, NADH (50 μM), lactate (10 mM), lactate dehydrogenase(20 U), pyruvate oxidase (2 U), peroxidase (15 U), flavin adeninedinucleotide (0.01 mM), cocarboxylase (0.2 mM), 4-aminoantipyrine (0.5mM), and N-ethyl-N-(2-hydroxy-3-sulfopropyl)-m-toluidine (0.02% w/v) ina total volume of 1 mL at pH 6.2. Coupling enzymes and reagents (that islactate, lactate dehydrogenase, pyruvate oxidase, flavin adeninedinucleotide, and cocarboxylase) allowed NADH oxidation to be coupled toH₂O₂ formation (pyruvate oxidation). In some assays (where specified),lactate and lactate dehydrogenase were replaced with alanine (10 mM) andalanine dehydrogenase (2 U). Assays were monitoredspectrophotometrically at 550 nm (6550 of 36.88 mM⁻¹ cm⁻¹ forquinoneimine dye product) over time in a thermostat-controlled cellholder at 30° C. as described above. Assays were pre-incubated for 2min. at 30° C. with all assay components except acetone. Assays wereinitiated by addition of acetone (2.5 mM).

Primary alcohol dehydrogenase coupled to primary alcohol oxidasesubstrate recycling assays. Assays were performed in 2 ml (1 cm pathlength) quartz cuvettes and contained potassium phosphate (25 mM),alcohol dehydrogenase (1 U), NADH (100 μM), alcohol oxidase (2 U),peroxidase (15 U), 4-aminoantipyrine (0.5 mM), andN-ethyl-N-(2-hydroxy-3-sulfopropyl)-m-toluidine (0.02% w/v) in a totalvolume of 1 mL at pH 6.2. Assays were monitored spectrophotometricallyat 550 nm (6550 of 36.88 mM⁻¹ cm⁻¹ for quinoneimine dye product) or at340 nm (ε₃₄₀ of 6.22 mM⁻¹ cm⁻¹ for NADH) over time in athermostat-controlled cell holder at 30° C. as described above. Assayswere pre-incubated for 2 min. at 30° C. with all assay components exceptethanol. Assays were initiated by addition of ethanol (50 μM or 5 μM).

Stability studies. A sufficient quantity of enzyme for each individualactivity assay (for example, 1.5 μg S-ADH) was aliquoted into 1.5 mLmicrocentrifuge tubes with specified concentrations of additives (forexample trehalose (10% w/v)) in buffer (25 mM MOPS, pH 7.6) and frozenat −80° C. for 1 h. Samples were then placed in a shelf freeze dryer(Virtis model Advantage ES) and held at −50° C. (shelf temperature) for16 h, and then increased to 20° C. for 4 h. Freeze-dried samples wereremoved and allowed to sit at room temperature (17 to 24° C.) over time.At specified time points, samples were re-hydrated and assayed asdescribed above.

Protein characterizations. Sodium dodecyl sulfate-polyacrylamide gelelectrophoresis (SDS-PAGE) was performed following the Laemmli procedure(Laemmli, U.K., Nature, 227:680-685 (1970)) using a 12% T, 2.7% C gel.“% T” indicates weight percent of total monomers, a measure of totalmonomer concentration, which is given by % T=100×((gramsacrylamide)+(grams cross-linker))/total gel volume (in mL); “% C”indicates weight percent of cross-linker, which is given by %C=100×grams cross-linker)/((grams acrylamide)+(grams cross-linker)); andthe cross-linker used was N,N′-methylene-bis-acrylamide. Electrophoresedproteins were visualized by staining with Coomassie Blue (PhastGel BlueR, catalog number 17-0518-01, Amersham Pharmacia Biotech). The apparentmolecular masses of polypeptides based on SDS-PAGE migration weredetermined by comparison with R_(f) values of standard proteins.N-terminal sequencing was performed by Commonwealth Biotechnologies,Inc. (Richmond, Va.). Protein concentrations were determined by using amodified biuret assay (V. J. Chromy et al., Clin. Chem, 20:1362-63(1974) with with δ-globulin as the standard.

Mass spectrometry analysis of enriched S-ADH and generation of peptideamino acid sequences was performed as follows. The S-ADH soluble proteinwas characterized by high-resolution two-dimensional gelelectrophoresis. Proteins (30 .μg) were solubilized for isoelectricfocusing (IEF) analysis in rehydration sample buffer consisting of 5 Murea, 2 M thiourea, 2% (w/v) CHAPS(3-[(3-cholamidopropyl)dimethylammonio-]-1-propanesulfonate), 2% (w/v)SB 3-10 (2-(decyldimethylammonio)propanesu-lfonate), 40 mM TRIS, 2 mMtributyl phosphine (added to rehydration solution just before use), and0.2% Bio-Lyte 3/10 (Bio-Rad, Hercules, Calif., cat. no. 163-2104).Protein/rehydration solution was rehydrated into 11 cm IPG ReadyStrip pH3-10 (Bio-Rad, Cat. no.163-2014) under passive conditions 0 volts, 20°C., 16 hrs.

One-dimensional isoelectric focusing was carried out on a Protean IEFcell (Bio-Rad, model no. 526BR02142) for 35,000 volt-hours using IPGReadyStrips (Bio-Rad). Following first dimension electrophoresis, gelswere equilibrated for 20 minutes in a buffer containing 20% glycerol,0.375 M Tris, 6 M urea, 2% SDS, and 5 M tributyl phosphine. IPGReadyStrips were placed on top of a Criterion™ precast 1 mm 4-20%gradient Tris-HCl-SDS gel (Bio-Rad, cat. no. 345-0036) and 0.5% warmAgarose containing 0.01% bromophenol blue (Bio-Rad, cat. no. 161-0404)was added to the remaining well. Electrophoresis was carried out on aCriterion mini electrophoresis cell (Bio-Rad, cat. no. 165-6001) at roomtemperature. The electrophoresis running buffer was prepared from a 10×Tris-glycine-SDS solution (Bio-Rad, cat. no. 161-0732). Followingassembly of the gel system and addition of the running buffer, theelectrophoresis was carried out at an initial current of 2 mA, 3500volts, 45 watts, for 1.5 hrs. The current was ramped up to 5 mA for 30minutes followed by 10 mA for 2-3 hrs. Typical run times were between4-5 hrs. Following electrophoresis, gels were stained in a bufferconsisting of 17% ammonium sulfate, 30% methanol, 3% phosphoric acid,and 0.1% coomassie brilliant blue G250 (Bio-Rad, cat no. 161-0436), forat least 12 hrs. Gels were rinsed with water and stored in 2% aceticacid until further processing.

Colloidal Coomassie-stained gel images were captured using Bio-RadsFluor-S MultiImager (Bio-Rad, cat. no.170-7700). Digital filteringalgorithms were used to remove non-uniform background without removingcritical image data. Internal standards (molecular weight markers) wereused initially to determine the molecular weight of the targetedproteins of interest. The molecular weight and pl of the S-ADH proteinwere determined by comparison of its position on the two-dimensional gelrelative to the protein standards.

Protein spots relative to S-ADH from the 2-D gel were excised manually.The gel pieces were macerated and destained with 25 mM ammoniumbicarbonate/50% acetonitrile in a 1.5 mL microfuge tube with vigorousshaking for 30 minutes. The blue-tinted destaining solution was removedand discarded with a fine-tip pipette. The destaining step was repeateduntil the stain was removed from the gel pieces. The gel pieces weredried under vacuum for 10 to 15 minutes. Proteins were digestedovernight at 37° C. in a total volume of 25 μL of sequence-grade,modified trypsin (Roche Diagnostics, Indianapolis, Ind.) at a finalprotein of 25 ng/μL in 25 mM ammonium bicarbonate. Peptides were elutedwith 50% acetonitrile and 0.5% trifluoroacetic acid. All peptide sampleswere concentrated, desalted, and detergents removed by using C18reversed-phase ZipTip™ pipette tips as described by the manufacturer(Millipore, Bedford, Mass., cat. no. ZTC18SO96).

The resulting tryptic peptides were analyzed directly by massspectrometry. Mass spectrometry experiments were carried out on aPerSeptive Biosystems (Framingham, Mass.) Voyager DE-STR equipped with aN₂ laser (337 nm, 3-nsec pulse width, 20-Hz repetition rate). The massspectra were acquired in the reflectron mode with delayed extraction.Internal mass calibration was performed with low-mass peptide standards,and mass-measurement accuracy was typically ±0.1 Da. All peptide sampleswere diluted in alpha.-cyano-4-hydroxycinnamic acid, which had beenprepared by dissolving 10 mg in 1 mL of aqueous 50% acetonitrilecontaining 0.1% trifluoroacetic acid.

Several tryptic peptide masses from S-ADH were further sequenced by oneof the following approaches by mass spectrometry as described below.

Approach 1: Tryptic digests of the protein were derivatized withchlorosulfonylacetyl chloride reagents as described by Keough T., LaceyM. P., Youngquist R. S. Proc. Natl. Acad. Sci. USA 1999; 96 7131. Thesulfonated sample was acidified with trifluoroacetic acid and cleaned updirectly using C18 mini-columns (ZipTips.™., Millipore). The derivatizedpeptides were eluted into α-cyano-4-hydroxycinnamic acid (Fluka, cat.no. 28480) and plated directly onto MALDI plates. Derivatized peptideswere analyzed on an Applied Biosystems Voyager DE-STR time-of-flightmass spectrometer equipped with a N₂ laser. All mass spectra wereacquired in the reflectron mode with delayed extraction. External masscalibration was performed with low-mass peptide standards, and massmeasurement accuracy was typically ±0.2 Da. PSD fragment ion spectrawere obtained after isolation of the appropriate derivatized precursorions using timed ion selection. Fragment ions were refocused onto thefinal detector by stepping the voltage applied to the reflectron in thefollowing ratios: 1.0000 (precursor ion segment), 0.9000, 0.7500,0.5625, 0.4218, 0.3164, and 0.2373 (fragment ion segments). Theindividual segments were stitched together using software developed byApplied Biosystems. All precursor ion segments were acquired at lowlaser power (variable attenuator=1980) for 100 laser pulses to avoiddetector saturation. The laser power was increased (variableattenuator=2365) for the remaining segments of the PSD acquisitions. ThePSD data were acquired at a digitization rate of 20 MHz; therefore, allfragment ions were measured as chemically averaged and not monoisotopicmasses.

Approach 2: Sequence tags were obtained from S-ADH tryptic peptides.Post source decay (PSD) fragment ion spectra were acquired for fourpeptides after isolation of the appropriate precursor ion by using timedion selection. Fragment ions were refocused onto the final detector bystepping the voltage applied to the reflector in the following ratios:1.0000 (precursor ion segment), 0.9000, 0.7500, 0.5625, 0.4218, 0.3164,and 0.2373 (fragment segments). The individual segments were stitchedtogether by using software provided by PerSeptive Biosystems. Allprecursor ion segments were acquired at low laser power (variableattenuator=1,450) for <256 laser pulses to avoid saturating thedetector. The laser power was increased for all of the remainingsegments of the PSD acquisitions. Typically, 200 laser pulses wereacquired for each fragment-ion segment. The PSD data were acquired at adigitization rate of 20 MHz. Mass calibration was performed with peptidestandards. Metastable decompositions were measured in all PSD massspectrometry experiments.

Approach 3: Sequence tags were obtained from S-ADH tryptic peptides byESI MS/MS the mass spectra were acquired on a Micromass Q-TOF2quadrupole/time of flight MS system.

Example 3

Initial electrochemical measurement of NADH and correlation tospectrophotometric data. 10 micron disc carbon fiber microelectrodeswere purchased (from Bioanalytical Systems (“BAS”), West Lafayette,Indiana (part number MF-2007)) and pretreated using the method of Kuhret. al. (63). The electrode surface was polished for 10 min. with 1 μmdiamond paste (Bioanalytical Systems) and sonicated in hot toluene for 2min. To remove residual polishing material, the microelectrode wasrinsed once in methanol and once in water, then sonicated twice in waterfor 1 min. The polished microelectrode was subsequently pretreatedelectrochemically in 1 M HCl by twice applying 10 cycles of 100 V/s from−200 mV to +1800 mV. Then the microelectrode was treated in 100 mMpotassium phosphate buffer by twice applying 10 cycles of 0 to +1200 mVat 100 mV/s. Background scans were then obtained from phosphate bufferalone. All potentials were referenced versus a Ag/AgCl referenceelectrode (Bioanalytical Systems). After baseline fast-scan cyclicvoltammograms (CVs) were obtained for the enzyme (1 U/mL) and NAD(P)H (2mM), the required volume of aqueous acetone was added (20 mM finalconcentration). The solution was quickly mixed, and fast-scan CVs wereobtained every 1 min. for 25 min. The buffer-only background wassubtracted from each CV with BAS 100W electrochemical software version2.3 (obtained from Bioanalytical Systems, West Lafayette, Ind.,hereinafter “BAS”).

Unless otherwise indicated, all electrochemical measurements wereperformed using a Bioanalytical Systems (BAS) Model 100A or Belectrochemical analyzer coupled to a BAS PA-1 preamplifier and aFaraday cage (part number MF-2500), wherein all waveforms were generatedand currents acquired via BAS 100W electrochemical software version 2.3.The data were processed using Microsoft Excel 97 SR-2 and BOMEM GRAMS/32version 4.04, Level II (Galactic Industries Corporation). Theelectrochemical cell was a custom-built 0.20 mL cell, constructed fromPlexiglas (acrylic polymer sheet, from Atofina Corp., Paris, France),containing a Ag/AgCl reference electrode, the pretreated carbon fibermicroelectrode, and a Pt wire auxiliary electrode.

To correlate spectrophotometric with electrochemical data for bothenzymes, the same reaction conditions were used for both analyses. ForS-ADH from T. brockii, the 1 mL reaction volume comprised finalconcentrations of 2 mM NADPH, 20 mM acetone, and 1 U S-ADH. For S-ADHfrom X. autotrophicus Py2, the 1 mL reaction volume comprised finalconcentrations of 2 mM NADH, 20 mM acetone, and 1 U S-ADH. For bothreactions, baseline A₃₄₀ was obtained for the enzyme and NAD(P)H versusa phosphate buffer blank. The cuvette containing the reaction solutionwas then removed from the spectrophotometer, and the 0.4 mL of solutionwas removed from the cuvette and combined with the remaining 0.6 mL. Therequired volume of aqueous acetone was added to the 1.0 mL reaction. Thesolution was mixed, 0.4 mL was added to the cuvette, and the cuvettereplaced in the spectrophotometer. The decrease in A₃₄₀ was thenmonitored for 30 min. using a Shimadzu UV-VIS-NIR scanningspectrophotometer (model UV-3101PC, Colombia, Md.). Data were acquiredusing UVPC Personal Spectroscopy Software version 3.9 (Shimadzu,Colombia, Md.) and processed using Microsoft Excel 97 SR-2. Quartzcuvettes with a 1 mm pathlength and a 0.4 mL volume were purchased (fromFisher Scientific, Pittsburgh, Pa., part number 14-385-906A).

Electrochemical measurement of acetone-dependent NADH consumption usingMeldola's Blue-modified carbon electrodes. A glassy carbon diskelectrode modified with the electrocatalyst Meldola's blue was preparedas follows. A 3-mm diameter glassy carbon electrode (BAS part numberMF-2012) was first wet-polished with a 1 m diamond suspension, sonicatedin deionized water for one minute, and then further polished with 0.05 malumina polishing suspension. The freshly polished electrode was washedthoroughly by sonication in deionized water and subsequently pretreatedelectrochemically in 5 mL deoxygenated 100 mM phosphate buffer (pH 7.2)by applying 20 cycles of 5 V/s from −500 mV to +300 mV, four times.After the cycling, a constant polarizing potential at −0.5 V was appliedfor 60 s. The electrochemically pretreated electrode was then soaked in0.5% of Meldola's blue (Aldrich, Milwaukee, Wis., catalog number32,432-9) at room temperature for 30 min. The electrode was rinsed withdeionized water before use.

Screen-printed carbon electrodes formulated with Meldola's Blue mediatorwere purchased from Gwent Electronic Materials Ltd. (Pontypool, UnitedKingdom). The disposable strips were configured in the geometrydescribed by Hart et al. and consisted of two screen-printed electrodesdeposited onto a polyethylene substrate. The working electrode wasgraphite carbon containing the electrocatalyst Meldola's Blue (partnumber C70902D2 from Gwent), and the reference/counter electrode wasAg/AgCl printed ink (part number C61003D7 from Gwent). The workingelectrode area was defined by printing an additional dielectric coating(part number D2000222D2 from Gwent). The electrode geometric area is 3×3mm, or 9 mm². The electrodes were pre-soaked in phosphate buffer for 10minutes before use to remove loosely bound Meldola's Blue.

The acetone-dependent consumption of NADH catalyzed by S-ADH wasmeasured with Meldola's blue-carbon electrodes prepared as above usingchronoamperometry in a 1 mL reaction volume containing 100 mM potassiumphosphate buffer (pH 7.2), NADH (500 μM), S-ADH (1 U), and varyingconcentrations of acetone. After a 2 min. incubation period, thepotential was stepped from open circuit to 68 mV (vs. Ag/AgCl) and thecurrent was recorded after 120 s.

Measurement of acetone-dependent consumption of NADH using commercialblood glucose disposable test strips. Disposable glucose biosensorstrips and reader (Precision Xtra Advanced Diabetes Management System)are available from MediSense (a division of Abbott Laboratories,Bedford, Mass.). 1 mL reaction volumes containing 25 mM potassiumphosphate buffer (pH 6.2), NADH (2 mM), S-ADH (20 U), and acetone (0.5,1.0, 1.5, 2.0 mM respectively) were incubated at room temperature. After5 min., a 20-μL aliquot was removed from each reaction mix and appliedto a disposable strip pre-inserted in the glucose meter. The meterreading value (mg/dL of glucose equivalent) was recorded and plotted tothe amount of acetone added.

Secondary alcohol dehydrogenase coupled to H₂O₂ formationelectrochemical assay. A disk platinum electrode (BAS part numberMF-2013) was used to monitor H₂O₂ produced by the S-ADH coupledenzymatic reaction in response to acetone concentration. Beforemeasurements the electrode surface was polished using Al₂O₃ paste for 1min. and then rinsed with deionized water, sonicated for 1 min. andrinsed with water again. The polished Pt electrode was then pretreatedelectrochemically by applying 10 cycles of 100 mV/s from +200 mV to +900mV. All potentials were referenced versus a Ag/AgCl electrode (BAS partnumber MF-2078). Assays contained potassium phosphate (100 mM, pH 7.2),purified S-ADH (1 U/mL), NADH (20 μM), lactate (100 mM), lactatedehydrogenase (5 U/mL), pyruvate oxidase (4 U/mL), flavin adeninedinucleotide (0.01 mM), cocarboxylase (0.2 mM), in a total volume of 0.5mL. Assays were initiated by addition of acetone. After a 2 min.incubation period, the potential was stepped from open circuit to 350mV. The oxidative current was recorded after 120 s and plotted againstacetone concentration.

Disposable electrode materials were evaluated to monitoracetone-dependent H₂O₂ produced by the coupled enzyme reaction using theidentical enzyme reagent system and similar electrochemical technique asdescribed above for the disk platinum electrode. Screen-printedplatinized carbon/graphite electrodes and cobalt phthalocyanine carbonelectrodes were purchased (part numbers C2000511D1, and C40511D8,respectively, Gwent Electronics Materials, Ltd.) with the same electrodegeometry as described earlier for the Meldola's Blue screen-printedcarbon electrodes. Screen-printed platinized carbon electrodes werepre-soaked in phosphate buffer for 5 min. before use. Assays wereinitiated by addition of acetone and incubated for 2 min. at which timethe potential was stepped from open circuit to 350 mV. The oxidativecurrent was recorded after 120 s. Cobalt phthalocyanine-modifiedscreen-printed carbon electrodes were pre-soaked in phosphate buffer for5 min. before use. After each addition of acetone, the reaction wasallowed to incubate for 3.5 min. Chronoamperometric measurements weremade with an initial quiet time of 5 s at 150 mV, and then the potentialwas stepped to 650 mV for 30 s and the current recorded. One cobaltphthalocyanine-modified screen-printed electrode was used for eachexperiment and then discarded.

A prototype disposable platinized carbon electrode was constructed bycutting ⅛ inch (3.06 mm) diameter circular disks (using a manual holepuncher) of Toray carbon paper (porous carbon paper) or cloth, loadedwith 20% (w/w) platinum nanoparticles (these platinum particles arenanonoparticles deposited on carbon; the platinum nanoparticle-loadedpaper or cloth was purchased from ETEK Division of De Nora NorthAmerica, Somerset, N.J., part number SLS-SPEC) and attached to ascreen-printed carbon working electrode (part number C10903D14 fromGwent Electronics Materials, Ltd.) using double-sided carbon tape (also⅛ inch (3.06 mm) diameter disk). In some experiments, 20 μM of non-ionicdetergent TRITON X-100 (t-octylphenoxypolyethoxyethanol; catalog numberT-8787, from Sigma Chemical Co.) or BRIJ 30 (tetraethylene glycolmonododecyl ether; catalog no. P-1254, from Sigma) was applied to theETEK material disk and allowed to dry before use. Before measurements,the electrode was pretreated electrochemically by applying 10 cycles of100 mV/s from +200 mV to +900 mV twice. Assays were initiated byaddition of acetone and incubated for 2 min. Chronoamperometricmeasurements were made with a quiet time of 2 s at 215 mV, and then thepotential was stepped from 215 mV to 350 mV vs. Ag/AgCl. The oxidativecurrent was recorded after 30 s.

Reflectance photometry measurement of acetone-dependent H₂O₂ formationusing glucose disposable test strips and correlation to electrochemicaldata. Disposable glucose biosensor strips and reader were purchased(OneTouch Basic read and strips from Lifescan, Inc., Milpitas, Calif.).Successive additions of 100 μM acetone were added to a 1 mL reactionvolume containing the S-ADH coupled enzyme system (as described above)and incubated at room temperature. Each acetone addition was allowed toreact for 4 min. and then a 20-μL aliquot was removed from the reactionmix and applied to a disposable strip pre-inserted in the glucose meter.The meter reading value (mg/dL of glucose equivalent) was recorded andplotted against the total concentration of acetone. H₂O₂ concentrationwas also monitored chronoamperometrically using a disk platinumelectrode as described above. The correlation between theelectrochemical assay and the colorimetric readings were plotted.

Enzyme-based electrochemical measurement of gas phase acetone. Gas phasesamples (0-10 ppm v/v) of acetone were prepared by injecting standardconcentrations of acetone into a calibrated airbag (10 L bag, CalibratedInstruments, Inc, Ardsley, N.Y.) filled with 7 L of water-saturated airand 1 L of dry air, and allowed to evaporate at 37° C. (about 30 min.).The gas samples produced from this system closely simulate human breathin terms of temperature and moisture content. The gas sampling systemwas calibrated (that is, concentration of acetone gas phase and liquidphase samples) using gas chromatography with a Hewlett Packard 5890 gaschromatograph equipped with flame ionization detection and an on-columninjector. 1 μL aqueous samples were applied to a 15 m long, coiledcapillary column (Nukol, 0.53 mm diameter with 0.50-μm layer of liquidphase, catalog number 25326, available from Supelco, Inc., Bellafonte,Pa.). The oven temperature was held at 40° C. for 4 min., then increasedat 25° C./min. to 200° C. The carrier gas flow rate was 5 mL/min. ofhelium.

Two types of sampling techniques were used to partition acetone from thegas phase into the liquid phase; a foam system, and a thin-aqueous layersystem. For the foam system, a piece of polyurethane foam was cut into acylindrical shape (19 mm long and 10 mm in diameter) so that the volumewas about 1 mL. The foam was boiled in water for 20 min. and theninserted into a 3 cc disposable plastic syringe. The syringe plunger wasinserted and pushed firmly to remove excess water and then removed.Before introducing gas phase acetone samples, 50 μL of water orphosphate buffer was loaded into the foam. Once the water contactedfoam, the surface tension sucked water into the foam cell and the waterdistributed evenly onto foam surface. The syringe containing wetted foamwas then connected via tubing to the gas sampling system and the gassample passed through the foam with a flow rate 5 L/min. for 12 secondseither by running a diaphragm pump or by manually pushing the airbag.This allowed the total gas sample volume to equal 1 L. After sampling,the syringe containing foam was quickly disconnected and the plungerre-inserted. The liquid was then squeezed out into an electrochemicalcell for electrochemical analysis or into a vial insert for gaschromatography analysis. For electrochemical measurements, theacetone-partitioned water sample was mixed with concentrated enzymesolution (S-ADH and coupling enzymes as discussed above) to make thedesired final enzyme solutions and incubated for 2 min. Theacetone-dependent H₂O₂ formed from the enzyme reaction was measuredchronoamperometrically as described above.

For the thin aqueous layer sampling method, the gas was released fromthe airbag in a fine stream at a flow rate of 500 mL/min. for 2 min. sothat the total volume of gas was equal to 1.0 L. In this experiment, theworking electrode was inverted (electrode surface facing up), so that asmall amount of enzyme solution (50 μL) forms a relatively thin layer ofliquid to cover the electrode surface. The gas was blown perpendicularto the liquid surface. The gas stream stirred the liquid to enhance themass transfer of acetone from gas phase into liquid phase. After the gassample flow, the enzyme solution was allowed to react for 1 min. Theacetone-dependent H₂O₂ formed from the enzyme reaction was measuredchronoamperometrically as described above. The current responses wereplotted against the gas-phase acetone concentration in the airbag.

1. A hand-held medical apparatus comprising: a. a housing; b. an inletfor receiving a sample of user breath; c. a sensor for detecting apre-determined breath component of said user breath and producing abreath-component signal over a measurement time; d. a sensing electricalcircuit in electrical communication with said sensor for sensing saidbreath-component signal, wherein the magnitude of said breath-componentsignal is a function of the concentration of said pre-determined breathcomponent in said breath sample to be received into said inlet; e. ananalog to digital converter in electrical communication with saidsensing electrical circuit for converting said breath-component signalto a digital signal; f. a microprocessor for processing said digitalsignal into at least one of a data signal and a user fat metabolismindicator; and g. a display in electrical communication with saidmicroprocessor for displaying said user fat metabolism indicator.
 2. Thehand-held medical apparatus of claim 1, wherein said sensor is anelectrochemical biosensor.
 3. The hand-held medical apparatus of claim2, further comprising means for removably retaining said electrochemicalbiosensor therewithin.
 4. The hand-held medical apparatus of claim 1,wherein said sensor comprises: a. means for removably receiving adisposable test matrix comprising an enzyme that selectively targets thepre-determined breath component as a substrate to produce a coloredproduct, wherein the amount of said colored product produced is afunction of the concentration of said pre-determined breath component insaid breath sample introduced into said inlet; b. a light source toilluminate said disposable test matrix; and c. a light detector todetect light reflected from said disposable test matrix and to createsaid breath-component signal.
 5. The hand-held medical apparatus ofclaim 1, wherein said sensor comprises: a. means for removably receivinga disposable test matrix comprising an enzyme that selectively targetsthe pre-determined breath component as a substrate to produce aluminescent product, wherein the amount of said luminescent productproduced is a function of the concentration of said pre-determinedbreath component in said breath sample introduced into said inlet; andb. a light detector to detect light emitted from said disposable testmatrix and to create said breath-component signal.
 6. The hand-heldmedical apparatus of claim 5, further comprising a light source toilluminate said disposable test matrix.
 7. The hand-held medicalapparatus of claim 5, wherein said disposable test matrix furthercomprises a working electrode, a counter electrode and a referenceelectrode.
 8. The hand-held medical apparatus of claim 1, wherein saidsensor is a thermosensor system.
 9. The hand-held medical apparatus ofclaim 8, wherein said thermosensor comprises a reference thermosensorand working thermosensor.
 10. The hand-held medical apparatus of claim1, further comprising means for receiving a compressible, porousmaterial wetted with a liquid and retaining it adjacent to saiddisposable electrode or said disposable test matrix, such that saidpre-determined breath component in said breath sample can partition intosaid liquid and then said liquid can be transferred to said disposableelectrode or said disposable test matrix by compressing saidcompressible, porous material.
 11. The hand-held medical apparatus ofclaim 1, further comprising data storage means.
 12. The hand-heldmedical apparatus of claim 1, further comprising a personal dataassistant in electrical communication with said microprocessor.
 13. Thehand-held medical apparatus of claim 12, wherein said personal dataassistant further comprises a clock for associating the time at whichsaid breath-component signal is produced.
 14. The hand-held medicalapparatus of claim 12, wherein said personal data assistant furthercomprises user input means by which information may be inputted asuser-input data, wherein said user-input data is stored and used tocreate a user fat metabolism indicator.
 15. The hand-held medicalapparatus of claim 12, wherein said personal data assistant furthercomprises at least one of the following: a. outgoing communication meansby which data is transmittable to a computer external to said personaldata assistant; and b. incoming communication means by which informationfrom a computer is receivable by said personal data assistant device ascomputer input data.
 16. The hand held medical apparatus of claim 1,wherein said inlet comprises a removable mouthpiece.
 17. The hand-heldmedical apparatus of claim 1, wherein said microprocessor integratessaid digital as a function of said measurement time to generate saiddata signal.